Metal Complex - DNA Interactions

Metal Complex–DNA Interactions

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

Metal Complex–DNA Interactions Edited by NICK HADJILIADIS Department of Chemistry, University of Ioannina, Greece and EINAR SLETTEN Department of Chemistry, University of Bergen, Norway

A John Wiley & Sons, Ltd., Publication

This edition first published 2009 © 2009 Blackwell Publishing Ltd Registered office John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, United Kingdom For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com. The right of the author to be identified as the author of this work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. The publisher and the author make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of fitness for a particular purpose. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for every situation. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of experimental reagents, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each chemical, piece of equipment, reagent, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. The fact that an organization or Website is referred to in this work as a citation and/or a potential source of further information does not mean that the author or the publisher endorses the information the organization or Website may provide or recommendations it may make. Further, readers should be aware that Internet Websites listed in this work may have changed or disappeared between when this work was written and when it is read. No warranty may be created or extended by any promotional statements for this work. Neither the publisher nor the author shall be liable for any damages arising herefrom. Library of Congress Cataloging-in-Publication Data Metal complexes : DNA interactions / edited by Nick Hadjiliadis, Einar Sletten. p. ; cm. Includes bibliographical references and index. ISBN 978-1-4051-7629-3 (cloth : alk. paper) 1. DNA-ligand interactions. 2. organometallic compounds. I. Hadjiliadis, Nick D. II. Sletten, Einar. III. Title: DNA interactions. [DNLM: 1. organometallic Compounds–chemistry. 2. Organometallic Compounds– pharmacology. 3. Antineoplastic Agents–therapeutic use. 4. DNA–chemistry. 5. Metals–therapeutic use. 6. Neoplasms–drug therapy. QV 290 M5831 2009] QP624.74.M48 2009 572.8′64–dc22 2008049834 A catalogue record for this book is available from the British Library Set in 10/12 Times by SNP Best-set Typesetter Ltd., Hong Kong Printed and bound in Great Britain by CPI Antony Rowe, Ltd, Chippenham, Wiltshire

To our families

Contents

Preface List of Contributors Acknowledgements A 1

2

Page xiii xv xix

BASIC STRUCTURAL AND KINETIC ASPECTS

1

Sequence-Selective Binding of Transition Metal Complexes to DNA Einar Sletten and Nils Åge Frøystein 1.1 Introduction 1.2 Ab initio Calculations and Photo-Cleavage Studies 1.3 NMR Spectroscopic Studies of Metal Binding to DNA Oligonucleotides 1.4 Summary of Theoretical and Experimental Evidence for Sequence–Selective Binding to DNA 1.5 Sequence-Specific Groove Binding Abbreviations References

3

Thermodynamic Models of Metal Ion–DNA Interactions Vasil Bregadze, Eteri Gelagutashvili and Ketevan Tsakadze 2.1 Introduction 2.2 Interactions of Metal Ions with DNA 2.3 Model and Mechanisms of Metal-Induced Formation of Point Defects 2.4 Conclusions Acknowledgements Abbreviations References

3 6 9 22 22 26 26 31 31 33 43 48 49 49 50

viii

3

4

B 5

6

Contents

Metal Ion Coordination in G-Quadruplexes Janez Plavec 3.1 Introduction 3.2 Cation Coordination and Stability of G-Quadruplexes 3.3 Structure of Sequences Consisting of Human Telomere Repeats 3.4 G-Quadruplexes Adopted by Promoter Regions 3.5 Bimolecular G-Quadruplexes of Analogues of the Oxytricha Telomere Repeat 3.6 Coordination of Cations within d[(TG4T)4] 3.7 Coordination of Cations within d[(G4T4G4)2] 3.8 Cation Movement within G-Quadruplexes 3.9 Summary Acknowledgements References Supramolecular Chemistry of Metal–Nucleobase Complexes P. J. Sanz Miguel, P. Amo-Ochoa, O. Castillo, A. Houlton and F. Zamora 4.1 Introduction 4.2 Discrete Architectures 4.3 Infinite Architectures 4.4 Overview of Metal Coordination Sites and Base-Base Hydrogen Bonding 4.5 M-DNA 4.6 Conclusion Acknowledgements References MEDICAL APPLICATIONS Platinum Drugs, Nucleotides and DNA: The Role of Interligand Interactions Giovanni Natile and Francesco Cannito 5.1 Introduction 5.2 Internucleotide Interactions 5.3 Guanine cis Amine Interactions 5.4 Solid-State Structures of Dynamic Nucleotides 5.5 Conformer Distribution in Cisplatin Adducts of G Derivatives 5.6 Retro Models Applied to Adducts with Tethered Guanine Bases 5.7 Conclusions and Perspectives Acknowledgements References Role of DNA Repair in Antitumour Effects of Platinum Drugs Viktor Brabec and Jana Kašpárková 6.1 Introduction 6.2 Human DNA Repair Systems

55 55 59 61 64 67 69 73 78 80 81 81 95

95 98 108 121 124 127 127 128 133 135 135 145 149 150 151 156 163 164 164 175 175 177

Contents

Specific Binding of Repair Proteins to DNA Modified by Antitumour Platinum Compounds 6.4 Repair of DNA Damage by Antitumour Platinum Compounds 6.5 Implications for Design of New Antitumour Platinum Compounds Acknowledgements References

ix

6.3

7

8

9

Telomeres and Telomerase: Potential Targets for Platinum Complexes Isabelle Ourliac-Garnier, Razan Charif and Sophie Bombard 7.1 Function of Telomeres 7.2 Structure of Telomeres 7.3 Telomerase 7.4 G-Quadruplex Structures and Small Molecules 7.5 Cisplatin 7.6 Interaction of Cisplatin and Related Platinum Complexes with G-Quadruplex Structures 7.7 Interaction of Cisplatin and Related Platinum Complexes with Telomeric DNA Duplexes 7.8 Interaction of Cisplatin and Related Platinum Complexes with Telomerase 7.9 Conclusion Acknowledgements Abbreviations References Towards Photodynamic Therapy of Cancer with Platinum Group Metal Polyazine Complexes David F. Zigler and Karen J. Brewer 8.1 Introduction 8.2 Study of Photophysics: Towards PDT 8.3 Photochemical Reactions of Metal Complexes with DNA 8.4 Designing Transition Metal Polyazines for DNA Photomodification 8.5 Cell Studies with Metal Complexes 8.6 Conclusions and Future Directions Acknowledgements Abbreviations References Platinated Oligonucleotides: Synthesis and Applications for the Control of Gene Expression Vicente Marchán and Anna Grandas 9.1 Therapeutic Applications of Synthetic Oligonucleotides 9.2 Platination as a Tool to Enhance Biological Effects

188 193 200 201 201

209 209 210 212 213 216 217 223 223 225 226 226 226

235 235 239 255 257 263 266 267 268 269

273 273 276

x

Contents

9.3 Synthesis of Platinated Oligonucleotides 9.4 Use of Platinated Oligonucleotides for Duplex Crosslinking 9.5 Use of Platinated Oligonucleotides for Triplex Crosslinking References

279 289 294 295

Rhodium– and Tin–DNA Interactions and Applications Kenneth D. Camm and Patrick C. McGowan 10.1 Introduction 10.2 Metal–DNA Interactions 10.3 Rhodium–DNA Interactions 10.4 Tin–DNA Interactions References

301

C

DNA RECOGNITION: NUCLEASES AND SENSORS

317

11

Groove-Binding Ruthenium(II) Complexes as Probes of DNA Recognition Jayden A. Smith, J. Grant Collins and F. Richard Keene 11.1 Introduction 11.2 Mononuclear Complexes 11.3 Dinuclear Complexes 11.4 Potential Biological Significance Abbreviations of Ligands References

10

12

13

DNA Recognition and Binding by Peptide–Metal Complex Conjugates Alexandra Myari and Nick Hadjiliadis 12.1 Introduction 12.2 Transition Metal Complex–Peptide Conjugates 12.3 Metallointercalator–Metallopeptide Conjugates 12.4 A Critical Survey and Future Perspectives 12.5 Conclusion Abbreviations References Artificial Restriction Agents: Hydrolytic Agents for DNA Cleavage Fabrizio Mancin and Paolo Tecilla 13.1 Introduction 13.2 DNA Hydrolysis 13.3 Free Ions and Mononuclear Complexes 13.4 Bimetallic Complexes 13.5 Conjugation with DNA-affine Subunits 13.6 Conjugation with Sequence-Selective Elements 13.7 The ARCUT System 13.8 A Critical Survey and New Perspectives

301 302 302 308 311

319 319 320 324 338 339 340

347 347 349 358 359 362 362 363

369 369 370 373 377 380 384 385 386

Contents

13.9 Conclusions Acknowledgements References 14

15

New Metallo-DNAzymes: Fundamental Studies of Metal–DNA Interactions and Metal Sensing Applications Zehui Cao and Yi Lu 14.1 Introduction 14.2 Metal Ions as Important Cofactors of DNAzymes 14.3 Selection of DNAzymes Using in vitro Evolution 14.4 Understanding Nucleic Acid Enzyme–Metal Ion Interactions 14.5 DNAzymes as Metal Ion Sensors 14.6 Summary Acknowledgements References Two-Metal-Ion-Dependent Catalysis in Nucleic Acid Enzymes Wei Yang 15.1 Chemistry of Nucleic Acid Synthesis, Cleavage and Strand Transfer 15.2 All DNA and RNA Polymerases Require Two Mg2+ Ions for Catalysis 15.3 Nucleases That Require Two Mg2+ Ions in the Active Site 15.4 Advantages of Two-Metal-Ion Catalysis: Specificity and Versatility 15.5 Concluding Remarks Acknowledgements References

D TOXICOLOGICAL ASPECTS 16

17

Structural Studies on the Mercury II-Mediated T-T Base-Pair Using NMR Spectroscopy Yoshiyuki Tanaka and Akira Ono 16.1 The History of the MercuryII–Mediated Thymine-Thymine Base Pair 16.2 Crystallographic Studies On HgII–Nucleobase Complexes 16.3 UV, UVCD and Vibrational (Ir/Raman) Spectral Studies 16.4 NMR Spectral Studies 16.5 Relationship Between 15n Chemical Shifts and Chemical Bonds 16.6 Applications of the T-HgII-T Base Pair 16.7 Biological Relevance of T-HgII-T Base Pairs 16.8 Concluding Remarks References Chromium-Induced DNA Damage and Repair Laura G. Little and Kent D. Sugden 17.1 Introduction 17.2 Nucleobase Oxidation

xi

389 389 390 395 395 396 397 399 403 410 411 411 415

415 417 420 428 430 430 430 437 439

439 440 441 444 452 454 455 457 457 463 463 468

xii

Contents

17.3 Sugar Oxidation 17.4 Chromium–DNA Binding 17.5 Cr(VI)-Induced DNA Crosslinks 17.6 Conclusions References 18

Arsenic-Induced Carcinogenicity: New Insights in Molecular Mechanism Andrea Hartwig and Tanja Schwerdtle 18.1 Introduction 18.2 Carcinogenicity and Cocarcinogenicity 18.3 Metabolism of Inorganic Arsenic 18.4 Modes of Action 18.5 Conclusions and Research Needs Acknowledgements Abbreviations References

Index

474 477 480 482 483

491 491 492 492 494 499 499 502 503 511

Preface

Metal ions and metal complexes have long been recognized as critically important components of nucleic acid chemistry both in regulation of gene expression and as promising therapeutic agents. Understanding how metal complexes interact with DNA has become an active research area at the interface between chemistry, molecular biology and medicine. In this volume, we attempt to bring together topics that span the breadth of this large area of research. The book is divided into four parts presenting recent developments in the field: Part A, Basic Structural and Kinetic Aspects, contains four chapters focusing on sequence-selective metal binding to DNA, thermodynamic models, metal coordination in G-quadruplexes and supramolecular networks of metal-nucleobases. Part B, Medical Applications, contains six chapters with the main focus on anticancer platinum drugs. Among the themes are the following: the role of interligand interactions, DNA repair in antitumour effects of platinum drugs, telomers and teleomerase as potential targets, photodynamic therapy, and synthesis and applications of platinated oligonucleotides for control of gene expression. A new and exciting field presented in Part C, DNA Recognition: Nucleases and Sensors, describes probes for DNA recognition, artificial restriction agents, metallo-DNAzymes for metal sensing applications and metal-ion-dependent catalysis in nucleic acid enzymes. Part D, Toxicological Aspects, deals with structural studies of mercury–DNA interactions, chromium-induced DNA damage and repair and the effect of arsenic-induced carcinogenicity. It has not been possible to cover all the topics that might find a place in a comprehensive coverage of metal–DNA interactions. The field is rapidly growing, spurred by significant improvements in experimental techniques. The applications of state-of-the art NMR spectroscopy, electrophoretic techniques, atomic force microscopy (AFM), etc. have dramatically expanded our knowledge on metal–DNA interactions during the last decade. The book presents pertinent up-to-date information for research scientists involved in, for example, the development of new and less toxic anticancer metallodrugs, the development of chemical nucleases and

xiv Preface

sensors, and the field of environmental and toxicological chemistry. Each chapter is written by experts on the various topics, and we hope this edition will prove to be a valuable contribution to the scientific community. Professor Nick Hadjiliadis (Ioannina, Greece)

Professor Einar Sletten (Bergen, Norway)

List of Contributors

Pilar Amo-Ochoa, Departamento de Química Inorgánica, Universidad Autónoma de Madrid, 28049 Madrid, Spain Sophie Bombard, Laboratoire de Chimie et Biochimie, Université Rene Descartes, 45 rue des Saints-Pères, 75270 Paris Cedex 06, France Viktor Brabec, Academy of Sciences of the Czech Republic, Institute of Biophysics, Kralovopolska 135, CZ-61265 Brno, Czech Republic Vasil Bregadze, E. Andronikashvili Institute of Physics, Georgian Academy of Sciences, 6 Tamarashvili Str, 0177 Tbilisi, Georgia Karen J. Brewer, Department of Chemistry, Virginia Tech, Blacksburg, VA 240610212, USA Kenneth D. Camm, School of Chemistry, University of Leeds, Leeds, LS2 9JT Francesco Cannito, Consorzio Interuniversitario di Ricerca in Chimica dei Metalli nei Sistemi Biologici (CIRCMSB), Bari, Via C. Ulpiani 27, I-70126 Bari, Italy Zehui Cao, Department of Chemistry, University of Illinois at Urbana-Champaign, A322 Chemical & Life Sciences Lab, 600 South Mathews Ave, Urbana IL 61801, USA Oscar Castillo, Departamento de Química Inorgánica, Facultad de Ciencia y Tecnología, Universidad del País Vasco, Apartado 644, E-48080 Bilbao, Spain

xvi

List of Contributors

Razan Charif, Laboratoire de Chimie et Biochimie, Université Paris Descartes, 45 rue des Saints-Pères, 75270 Paris Cedex 06, France J. Grant Collins, School of Physical, Environmental and Mathematical Sciences, University of New South Wales Australian Defence Force Academy, Canberra, ACT 2600, Australia Nils Åge Frøystein, Department of Chemistry, University of Bergen, Allégt. 41, 5007 Bergen, Norway Eteri Gelagutashvili, E. Andronikashvili Institute of Physics, Georgian Academy of Sciences, 6 Tamarashvili Str, 0177 Tbilisi, Georgia Anna Grandas, Departaments de Química Orgànica (Facultat de Química), IBUB Universitat de Barcelona, Martí i Franquès 1-11, 08028 Barcelona, Spain Nick Hadjiliadis, Department of Chemistry, University of Ioannina, Ioannina 45110, Greece Andrea Hartwig, Institut für Lebensmitteltechnologie und Lebensmittelchemie, Der Technische Universität Berlin, Gustav-Meyer – Allee 25, 13355 Berlin, Germany Andrew Houlton, Chemical Nanoscience Laboratory, School of Natural Sciences, Newcastle University, Newcastle upon Tyne, NE1 7RU, UK Jana Kašpárková, Academy of Sciences of the Czech Republic, Institute of Biophysics, Kralovopolska 135, CZ-61265 Brno, Czech Republic F. Richard Keene, School of Pharmacy & Molecular Sciences, James Cook University, Townsville, Queensland 4811, Australia Laura G. Little, Department of Chemistry, The University of Montana, Missoula, Montana 59812, USA Yi Lu, Department of Chemistry, University of Illinois at Urbana-Champaign, A322 Chemical & Life Sciences Lab, 600 South Mathews Ave, Urbana IL 61801, USA Fabrizio Mancin, Dipartimento di Scienze Chimiche, Università de Padova, Via Marzolo 1, I-35131, Padova, Italy Vicente Marchan, Departaments de Química Orgànica (Facultat de Química), IBUB Universitat de Barcelona, Martí i Franquès 1-11, 08028 Barcelona, Spain Patrick C. McGowan, School of Chemistry, University of Leeds, Leeds, LS2 9JT Pablo J. Sanz Miguel, Departamento de Química Inorgánica, Universidad Autónoma de Madrid, 28049 Madrid, Spain

List of Contributors xvii

Alexandra Myari, Department of Chemistry, University of Ioannina, Ioannina 45110, Greece Giovanni Natile, Dipartimento Farmaco-Chimico, Università degli Studi di Bari, Via E.Orabona 4, 70125 Bari, Italy Akira Ono, Department of Material & Life Chemistry, Faculty of Engineering, Kanagawa University, 3-27-1 Rokkakubashi, Kanagawa-ku, Yokohama, Kanagawa 221-8686, Japan Isabelle Ourliac-Garnier, Institut Curie, Centre de Recherche, 26 rue d’Ulm, 75248 Paris, Cedex 05, France Janez Plavec, Slovenian NMR Center, National Institute of Chemistry, Hajdrihova 19, SI-1000 Ljubljana, Slovenia Tanja Schwerdtle, Institut für Lebensmitteltechnologie und Lebensmittelchemie, Der Technische Universität Berlin, Gustav-Meyer – Allégt 25, 13355 Berlin, Germany Einar Sletten, Department of Chemistry, University of Bergen, Allgt. 41, 5007 Bergen, Norway Jayden A. Smith, School of Pharmacy & Molecular Sciences, James Cook University, Townsville, Queensland 4811, Australia Kent D. Sugden, Department of Chemistry, The University of Montana, Missoula, Montana 59812, USA Yoshiyuki Tanaka, Laboratory of Molecular Transformation, Graduate School of Pharmaceutical Sciences, Tohoku University, Aobayama, Aoba-ku, Sendai, Miyagi 980-8578, Japan Paolo Tecilla, Dipartimento di Scienze Chimiche, Università di Trieste, via Giorgeri, 1-34127 Trieste, Italy Ketevan J. Tsakadze, E. Andronikashvili Institute of Physics, Georgian Academy of Sciences, 6 Tamarashvili Str, 0177 Tbilisi, Georgia Wei Yang, National Institute of Diabetes and Digestive and Kidney Diseases, NIH, Bethesda, Maryland, MD 20892 USA Felix Zamora, Departamento de Química Inorgánica, Universidad Autónoma de Madrid, 28049 Madrid, Spain David F. Zigler, Department of Chemistry, Virginia Tech, Blacksburg, VA 240610212, USA

Acknowledgements

The editors would like to thank all authors for their important and high quality contributions to this book. Great thanks go to Jorunn Sletten for invaluable assistance in the editing process. Finally we thank Wiley for guidance through the production of this book.

Figure 3.2 (Plate 1) Variation in strand stoichiometry and topology of inter- and intramolecular G-quadruplex structures: (a) tetramolecular structure with all four parallel strands and all anti glycosidic torsion angles; (b) bimolecular and (c) monomolecular structures with antiparallel strands and alternating syn (in orange) and anti (in blue) orientations across glycosidic bonds

Figure 3.4 (Plate 2) Polymorphism of four repeats of human telomeric sequence: (a) intramolecular G-quadruplex formed by d[AG3(T2AG3)3] as determined by NMR spectroscopy in the presence of Na+ ions in solution;92 (b) the G-quadruplex fold with parallel GGG strands and double-chain reversal loops as determined by X-ray crystallography for the same sequence crystallized in the presence of K+ ions;104 (c) the topology of the four-repeat sequence in K+ ion containing solution (so called Hybrid-1 structure);204,205,207 (d) the fold of major form of the unmodified four-repeat sequence in K+ ion containing solution (so called Hybrid-2 structure).210 For clarity, only G-bases and their syn (in orange) and anti (in blue) orientations across glycosidic bonds are shown Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

Figure 3.5 (Plate 3) Representative structures of gene promoter regions: (a) G-quadruplex fold formed by guanine tracts of c-myc-2345 and c-myc-1245 sequences originating from the c-myc promoter, determined by NMR;231 (b) topology adopted by the c-myc-23456 sequence containing five of the six guanine tracts of the c-myc promoter region;232 (c) NMR structure of unprecedented intramolecular G-quadruplex formed by a G-rich sequence in the c-kit promoter in K+ solution233

Figure 3.6 (Plate 4) Topology of analogues of the Oxytricha nova telomere repeat sequence: (a) the structure of bimolecular d[(G4T4G4)2] quadruplex was one of the first high resolution structures of G-quadruplexes determined;96 (b) unique folding topology of d[(G3T4G4)2] with (3 + 1) strand orientations;244 (c) topology of d[(G4T4G3)2];243 (d) asymmetric structure of d[(G3T4G3)2]238

Figure 3.8 (Plate 5) Cation coordination within G-quadruplex structures adopted by d(TG4T) as revealed by X-ray crystallography: (a) Na+ ions (pink balls) within the 0.95 Å resolution structure (PDB ID 352D);157 (b) Na+ ions in the structure crystallized using lithium sulfate as the main precipitating agent (PDB ID 2O4F);246 (c) arrangement involving both Ca2+ (green balls) and Na+ (pink balls) cations crystallized in a mixed ion environment (PDB ID 2GW0);139 (d) Tl+ (red balls) and Na+ (pink ball) ions within the 2.2 Å resolution structure (PDB ID 1S45);247 (e) Tl+ (red balls) and Na+ (pink balls) ions within the 2.5 Å resolution structure (PDB ID 1S47);247 (f) Tl+ (red balls) and Na+ (pink balls) ions within the G-quadruplex exhibiting a T-quartet at the 3′-end of the strands (shown at the top of this structure) (PDB ID 1S47).247 Individual strands are shown in a wire frame model, while guanine bases are presented in stick representation to demonstrate the extent of out of plane bending. Cations are shown as coloured balls. Na+ with the smallest ionic radius can vary in coordination geometry from being within the plane of a G-quartet to being equidistant between two adjacent Gquartets. The larger ions are exclusively coordinated between two adjacent quartets

Figure 3.9 (Plate 6) Localization of cations within bimolecular G-quadruplexes: (a) Na+ ions (pink balls) within the d[(G4T4G4)2] quadruplex in crystals of the Oxytricha nova telomere end-binding protein with telomere DNA (PDB ID 1JB7);158 (b) K+ ions (grey balls) within d[(G4T4G4)]2 as revealed by X-ray crystal structure at 2.0 Å resolution (PDB ID 1JRN);99 (c) K+ ions (grey balls) within d[(G4T4G4)2] as revealed by X-ray crystal structure at 1.5 Å resolution (PDB ID 1JPQ);99 (d) five Tl+ ions (red balls) within d[(G4T4G4)2] as revealed by X-ray crystal structure at 1.55 Å resolution (PDB ID 2HBN);129 (e) ammonium ions within d[(G4T4G4)2] as established by NMR measurements in solution;162 (f) two ammonium ions within d[(G3T4G4)2] established by NMR studies in solution.176 Individual strands are shown in wire frame model, while guanine bases are presented in stick representation to demonstrate the extent of out of plane bending. Na+ ion can be localized within the plane of a G-quartet or between two adjacent G-quartets. The larger K+, Tl+ and 15NH4+ ions are exclusively coordinated between two adjacent quartets

platinum

hydrogen

nitrogen

carbonium

oxygen

phosphorus

Figure 5.9 (Plate 7) X-ray structures of two cis-A2Pt(5′-GMP)2 adducts showing how in the DHT conformer (top, based on data from Ref. 117) the 5′-phosphate is directed towards the cis amine, while in the ΛHT conformer (bottom, based on data from Ref. 118) the 5′phosphate is directed towards the cis guanine. For simplicity only the chelate ring atoms of the carrier ligands (Me4dach in the former case and Me4en in the latter case) are shown

HH1

∆HT1

HH2

platinum

nitrogen oxygen

carbonium phosphorus

Figure 5.13 (Plate 8) Minimum energy models (from NMR-restrained MMD calculations) of the anti/anti HH1 L (top) and anti/syn DHT1 L (centre) forms of (S,R,R,S)-bip-Pt(d(GpG)) and the anti/anti HH2 R (bottom) form of (R,S,S,R)-bip-Pt(d(GpG)) (the conformation of the 5′-G is given first, then that of 3′-G; only the chelate-ring atoms of the bip ligand are shown). Based on data from Refs 101 and 106

view from the major groove

view from the minor groove

platinum

nitrogen oxygen

carbonium phosphorus

Figure 5.15 (Plate 9) Central portion (four base-pairs) of oligonucleotides containing G/G intrastrand (top) and interstrand (bottom) crosslinks (based on data from Refs 114 and 143, respectively)

A H H N

T N

5' HO

-

N

O

O O P O O

O N

N

O O P OO-

O O P OO

N

O

H

O

N

N

H

N

N

3'

3'

O

H

HN

O

OO P O O

N

N

O

-

H

O

major groove

OH

5'

NH

minor groove

C

N

G

Figure 8.2 (Plate 10) Complimentary pairs of nucleotides that make up double helix DNA (Inset, A = adenine, T = thymine, G = guanine, C = cytosine, DNA = deoxyribonucleic acid). Atoms are carbon (grey), hydrogen (white), nitrogen (blue), oxygen (red) and phosphorous (pink)

N

N RuII N

N N

N

Cl

Cl

N

RhIII N

N N

N

N RuII N

N N

N

MC = [{(bpy)2Ru(dpp)}2RhCl2]Cl5 Cells treated with MC

illumination area

growth

after 48 hr growth period

calcein AM imaging of live cells

ethidium imaging of dead cells

Figure 8.20 (Plate 11) Micrographs of Vero cells pretreated with [{(bpy)2Ru(dpp)}2RhCl2]C l5, rinsed, and illuminated with 400–1000 nm light showing the high level of light-activated cell killing [bpy = 2,2′-bipyridine, dpp = 2,3-bis(2-pyridyl)pyrazine]. (From left to right) immediately after photolysis (light exposure within circle); after 48 hour growth period; live cell (green) visualized with calcein AM fluorescent dye; dead cell (red) visualized with ethidium homodimer-1 fluorescent dye84

Figure 14.2 (Plate 12)

Schematic representation of the in vitro selection process

Figure 14.5 (Plate 13) Single molecule FRET study of the 8-17 DNAzyme: (a) time traces of the FRET signals and changes upon injection of 100 mM Zn2+ at 21 s; (b) distribution of single DNAzyme molecules with different FRET efficiencies before and after injection of 20 mM Pb2+; (c) proposed reaction pathways of the DNAzyme in the presence of Zn2+/Mg2+ and Pb2+. (Reprinted with permission from Nat. Chem. Biol., 2007, 3, 763–768.)

Figure 14.7 (Plate 14) Design and performance of DNAzyme-based UO22+ sensor. Competing metals were tested from 10 µM to 1 mM, and UO22+ (the last three bars) was tested from 0 to 10 nM. Inset: Sequence, schematics and fluorescent detection of the DNAzyme-based UO22+ sensor. (Juewen Liu et al., A catalytic beacon sensor for uranium with parts-per-trillion sensitivity and millionfold selectivity, Proc. Natl. Acad. Sci. USA, 104, 2056–2061. Copyright 2007, National Academy of Sciences, USA.)

Figure 15.3 (Plate 15) Three types of catalytic centres found in DNA and RNA polymerases; (A) Phage T7 DNA polymerase represents the pol I-like active centre, which is found in the A, B and Y-family DNA polymerases, retroviral reverse transcriptases, phage RNA polymerases and viral RNA-dependent RNA polymerases. The catalytic residues (highlighted in blue and red) include two Asps and one Glu on two adjacent antiparallel strands (highlighted in cyan); (B) Human DNA pol b represents the pol b-like catalytic centre, which is found in the C and X-family DNA polymerases, poly-rA polymerases and CCA-adding enzymes. The catalytic residues include three conserved Asps on two adjacent parallel strands; (C) S. cerevisiae RNA pol II represents all bacterial and eukaryotic DNA-dependent RNA polymerases. The catalytic core consists of a b-barrel with three catalytically essential Asps on a loop (highlighted in cyan). All of these polymerases contain two absolutely conserved Asps, which together with the a-phosphate of (d)NTP jointly coordinate the two Mg2+ (shown as green spheres)

RNase H

MutH

Figure 15.4 (Plate 16) Two families of nucleases that have been shown to use two-metal-ion catalysis: (A) RNase H is a prototype of the superfamily, which includes nucleases, retroviral integrases, RuvC Holliday junction resolvase and dicer argonaute. The two catalytically essential Asps are located on two adjacent parallel strands (highlighted in cyan). Two additional carboxylates may be required for catalysis. Two Mg2+ are jointly coordinated by the carboxylates and the scissile phosphate; (B) MutH represents another superfamily, which includes type IIP restriction endonucleases, T7 endonuclease I (a Holliday junction resolvasae) and T4 exonuclease I. The catalytic residues, Asp (D), Glu (E) and Lys (K) are located on the diverging point of a b-hairpin (highlighted in cyan). The loop that contains the first Asp, which plays the central role in Mg2+ coordination, is often disordered in the MutH-family nucleases in the absence of a properly aligned substrate or divalent cations. A diagram of the metal ion coordination in each case is shown below the actual structure

Figure 15.6 (Plate 17) Coordination of the two metal ions changes during RNA cleavage by RNase H: (A) Enzyme–substrate complex; (B) Enzyme–product complex

Figure 15.7 (Plate 18) Coordination of the two metal ions in Group I intron. The two metal ions have similar coordination geometry (hence symmetric) and can reverse their roles as ‘A’ and ‘B’ metal ions in the successive steps of RNA splicing reaction. (Reproduced from Yang, W. et al., Making and breaking nucleic acids: two-Mg2+-ion catalysis and substrate specificity, Mol. Cell, 2006, 22, 5–13)

Part A Basic Structural and Kinetic Aspects

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

1 Sequence-Selective Binding of Transition Metal Complexes to DNA Einar Sletten and Nils Åge Frøystein

1.1 Introduction The biological significance of the interaction between metal ions and nucleic acids has become a rather well-established fact. One may mention the observed necessity for the presence of metal ions in many natural processes where nucleic acids play the dominant role. The effect of platinum-based chemotherapeutic drugs probably originates from their attack on DNA. Another aspect of metals in biological systems is the increased flux of metals in the environment during the last decades. An assessment of the toxic effect of an unnatural metal ion concentration must include information on the processes in which the metal can participate. In a comparison of metal carcinogenicity in humans based on several experimental factors, Cr and Ni turned out to be the most potent carcinogens.1 The nucleic acid monomers, guanine (G), adenine (A), thymine (T) and cytosine (C) have different metal ion affinities. The order of stability of 3d transition metal ion–nucleobase complexes are: G > A, C > T.2 At physiological pH the preferred binding sites on the nucleobases are: guanine N7, adenine N1 and/or N7, cytosine N3, thymine O4. For nucleotides the relationship between phosphate and base binding is dependent on the type of metal ion. Eichhorn and Shin3 studied the effect of various metal ions on the melting temperature of DNA (Figure 1.1). The authors suggest that magnesium ions increase Tm by binding to phosphate and Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

4

Sequence-Selective Binding to DNA

Mg Co Ni

75

Mn

Zn

Tm °C

65

55 Cd

45

Cu 35 0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

MOLES M+2/DNA (P)

Figure 1.1 Variation of Tm of solutions of DNA as a function of divalent metal ion concentration. (Reprinted with permission from J. Am. Chem. Soc., 1968, 90, 7327. Copyright 1968 American Chemical Society.)

stabilizing the double helix, whereas copper ions decrease Tm by binding to the bases and destabilizing the double helix. Based on the metal-induced variation in Tm they suggested that the relative metal affinity to the phosphate backbone of DNA follows the order Mg2+ > Co2+ > Ni2+ > Mn2+ > Zn2+ > Cd2+ > Cu2+. This implies that the binding of an individual metal ion may involve phosphate and base on the same molecule or form a linkage between two different nucleotides. An example of the latter situation is the mononucleotide–metal ion binding pattern observed for the Cu-(GMP) complex, where Cu2+ ions are bridging the GMP ligands through alternating N7–Cu–phosphate bonds (Figure 1.2).4 When nucleobases are incorporated into a duplex DNA matrix, the affinities towards metal ions are modified. It has been shown that several divalent metal ions, like Mn2+, Cu2+ and Pt2+ prefer GC-rich regions, while Hg2+, for example prefer ATrich regions.5,6 A more detailed picture indicates that metal binding to base residues is sequence-dependent, i.e. not all guanines in a particular sequence show identical affinity towards a specific type of metal ion.7–10 As a consequence, one may envisage designing metal complexes that can bind selectively to chosen sequences of DNA.

Introduction

5

Figure 1.2 The nucleobase–metal–phosphate linkage in the crystal structure of Cu(5′GMP) 4

Such complexes may be used as drugs that block specific gene expression associated with a certain disease. Metal ions can interact with nucleic acids in two distinct modes of binding: diffuse binding and site binding, both of which are important for the structure and function of nucleic acids.11a,b In the diffuse binding mode the metal and the nucleic acid retain their hydration layer and the interaction is through water molecules.11c This is a long-range Coulombic interaction, in which positive metal ions accumulate around the nucleic acid in a delocalized manner; for example, the counterion atmosphere that all nucleic acids possess is made up of diffusely bound positive ions. In the site-binding mode the metal is coordinated to specific ligands on the nucleic acid; the coordination can either be direct (termed inner-sphere) or through a water molecule (termed outer-sphere). In the outer-sphere binding mode only the innermost hydration layer of the metal is kept intact, and the metal and the nucleic acid ligand(s) to which the metal is coordinated share solvation shells. In inner-shell binding there is direct contact between the metal and the nucleic acid. Dehydration of the metal ion and the nucleic acid binding site therefore has to occur before an inner-shell bond is formed. The mechanism of inner-sphere binding is likely to be initiated by a diffuse binding mode, in which the metal and the nucleic acid are separated by no more than two layers of solvent molecules.11d This step is diffusion controlled. The next step is that the metal ion and the nucleic acid form an outer-sphere complex, separated only by one layer of solvent molecules. This step primarily depends on electrostatic attractions and hydrogen bonding between the metal and the nucleic acid.12 In the final step the metal and the nucleic acid come into direct contact (innersphere binding). Here the nucleophilicity of the coordination site plays a crucial role. In the last two steps steric effects are also important. Several attempts have been made to quantify the importance of accessibility and molecular electrostatic potential (MEP) at the site where the inner-sphere complex/covalent bond is formed.13,14 In these studies a reasonable correlation between these two important

6

Sequence-Selective Binding to DNA

factors exists, and has been used to predict which DNA site is the most reactive to metalation or methylation. In this chapter we present data on sequence-selective interactions between metal complexes and nucleic acids. In the outline we will distinguish between (i) site-selective inner-sphere metal coordination of nucleobases, and (ii) the selectivity of fully hydrated species located in the minor or major groove through hydrogen bonding and electrostatic interaction. In the former case a further distinction will be made between labile and nonlabile metals.

1.2 Ab initio Calculations and Photo-Cleavage Studies The highest occupied molecular orbital (HOMO) of DNA nucleobases plays a crucial role in metal coordination by interacting with the lowest unoccupied molecular orbital (LUMO) of metal ions.15 The calculations of HOMOs of macromolecules such as duplex DNA are extremely difficult. Consequently, there has been little focus on the role of the HOMO/LUMO in studies of DNA–metal ion interactions. Theoretical calculations of DNA bases have mostly focused on ionization potentials (IP) of monomeric nucleobases and the stability of the nucleobase pair in neutral and radical cation states.16 About ten years ago the Saito group published the first extensive studies on the variation of nucleobase IPs and localization of HOMOs as a function of base stacking, using high-level ab initio calculations.17 The IPs of four base monomers and 16 sets of nearest-neighbour stacked nucleobases in the B form were calculated. It was found that the GG/CC system has the lowest IP among ten possible stacked nucleobase pairs and that approximately 70% of the HOMO is localized on the 5′-G of 5′-GG-3′. These calculations indicate that the 5′-G of 5′-GG3′ is the most electron-donating site in B-DNA. The origin of IP lowering as a result of base stacking was further investigated by calculations of HOMO energy distribution as a function of the twist angle of a GG dimer.17 Within a normal range of twist angles for B-DNA (−25 ° to −45 °), IP values of GG are between 7.2 and 7.4 eV and the HOMO is predominantly localized on 5′-G. This implies that in B-form DNA the 5′-side G of the 5′-GG-3′ sequence is the most strongly interacting site with electrophiles. This principle may be very important in governing sequence-selective metal binding to DNA. According to the theoretical calculations, HOMOs of GG sites in duplex DNA should serve as the most reactive one-electron oxidation sites. In order to verify this assumption experimentally, Saito et al. performed laser flash photolysis of duplex DNA oligonucleotides with added photosensitizer (photocleaving aminoacid, PCA) and subsequent hydrolysis dephosphorylation with alkaline phosphatase (Scheme 1.1).18 The G3 : G4 ratio (84 : 16) determined by HPLC implies that the major degradation pathway of the hexamer involves the G3 site – the most readily oxidizable site according to theoretical calculations. It should be noted that photo-irradiation of the single-strand alone results in a nonselective cleavage at G3 and G4 in the ratio 1 : 1.

Ab initio Calculations

5´- T1T2G3G4T5A6

7

(T1 T2 + G3TA) + (TTG4 + TA) + PCA

3´-A-A-C-C-A-T

3´-A-A-C-C-A-T Scheme 1.1

Figure 1.3 Distribution of HOMOs (normalized to the largest value) in B-form duplex 5-mers obtained by ab initio calculations. The arrows indicate the distribution of the HOMOs normalized to the largest HOMO as 100 20

Further work by the Saito group involved ab initio calculations of HOMOs for a wide variety of double-stranded G-containing 5-mer sequences with B-form geometry using GUSSIAN 9x1at the HF/6-31G* level.19,20 For the quantum mechanical studies, all of the sugar backbones of the 5-mers were removed from the coordinate file and replaced by methyl groups. A few examples of the distributions of HOMOs in the duplex 5-mers are shown in Figure 1.3. The general trend for HOMO distribution is that the HOMO of stacked GG doublets is localized overwhelmingly on the 5′-G, regardless of the 3′- and 5′flanking residues (A, C or T). Bearing in mind that the model used is rather crude (all of the sugar backbone replaced by methyl groups), further discussion of more subtle sequence-specific differences is not warranted. Further experimental support for the theoretical results was obtained by studying the oxidation of oligodeoxynucleotides (ODN) with Co2+ ions and benzoyl peroxide using PAGE analysis of the reaction mixture after hot piperidine treatment. Sequence-dependent G-cleavage was observed for double-stranded ODN, whereas nonselective equal G cleavage was observed for single-stranded ODN. The relative rates of sequence oxidation were determined by densitometric assay of the ODN cleavage bands. Experimentally observed relative rates of G oxidation matches well with the calculated HOMOs of the G-containing sequences, implying that the Co2+ ion is coordinated more strongly to the G having the larger HOMO.20 Comparable theoretical calculations by Senthikumar et al.21 on stacked XGY triplets with B-form geometries, including sugar and phosphate groups, show that the site energy is strongly influenced by the type of nucleobase at the 3′ position.

8

Sequence-Selective Binding to DNA

When C or T is present at the 3′ position, the site energy at the guanine was found to be up to 0.44 eV higher than for A or G at this position. The influence of the base at the 5′ position was much smaller, the variation in site energy being less than 0.1 eV. The amount of charge on a certain G was calculated from the coefficients of the guanine fragment orbitals. It was concluded that the neighbouring base at the 3′position to a large extent determines the charge distribution and therefore the oxidative damage on a sequence of guanine bases.21 Further work by the Saito group on experimental mapping of G-rich hot spots included photo-induced cleavage of double-stranded 32P-end-labelled oligodeoxynucleotides (ODNs) 30-mers possessing two different G-containing sequences (5′TXGYT-3′) and a 5′-TTGGT-3′ step as a standard on the same strand.22 Under low photo-irradiation conditions, only the cleavage bands of 5′-Gs of the two GG steps and the middle step of the GGG triplet were observed by hot piperidine treatment. Quantitative densitometric assay was used to determine the relative amounts of cleavage products. The experimental results were compared to IP values calculated for 16 sets of base-paired G- and GG-containing 5-mers. A plot of the log of the relative reactivity (krel) toward photo-induced one-electron oxidation versus calculated IP is shown in Figure 1.4. A different explanation for the enhanced reactivity of the 5′-G of a GG step compared to the 3′-G is presented by the Schuster group.23 Theoretical calculations of base-paired quartets, d(5′-XGGX-3′)/d(5′-YCCY-3′), suggested that electronic factors may not be the primary determinant of the reaction selectivity for GG steps. Instead the authors propose, based on molecular dynamic (MD) simulations on BDNA oligomers, d(5′-GXXGGXXG-3′)/d(5′-GYYGGYYG-3′), where X = A,T,U and Y is the complementary base, that ‘there is an important steric contribution to the preference for reaction at the 5′-G in the GG doublets’. Photo-cleavage experiments carried out on the series of B-DNA oligomers showed that for GG steps in

Figure 1.4 A plot of the log of the relative reactivity (krel) toward photo-induced one-electron oxidation versus calculated ionization potential (IP). The relative reactivity was obtained from densitometric assay of G bands with TGG as a standard (krel = 1.0) under single-hit conditions. (Reprinted with permission from J. Am. Chem. Soc., 1998, 120, 12687. Copyright 1998 American Chemical Society.)

NMR Spectroscopic Studies

9

the context AGGA, the ratio of 5′ to 3′ reactivity was 1.8 ± 0.1, and for GG in the context TGGT, the ratio was 6.1 ± 0.3. The authors propose that the accessibility of H2O to the reaction site determined by the steric blocking by the methyl group plays the dominant role for the observed sequence-selectivity, rather than electronic effects.23

1.3 NMR Spectroscopic Studies of Metal Binding to DNA Oligonucleotides 1.3.1 NMR Methodology Most early nuclear magnetic resonance (NMR) studies on DNA involved complementary homopolymers and self-complementary, alternating copolymers, e.g. poly(dA)/poly/dT).24 The development of efficient and rapid methods of large-scale oligonucleotide syntheses has made it possible to design heteropolymeric sequences of high purity. Dodecamer (12 base pair) sequences adopting a normal B-DNA double-helical conformation, are assumed to complete a full turn of a right-handed helix. The structure of such a mini-helix is probably sufficiently close to that of real DNA to serve as a realistic model for determining preferred metal binding sites. The effects of adding paramagnetic metal ions to an aqueous solution of DNA fragments may be monitored by observing the decrease in spin-lattice (T1) and spinspin (T2) relaxation times (related to line-broadening) for protons close to the metal centres. Paramagnetic metal ions may be classified according to their electronic correlation times, i.e. as relaxation probes producing broad lines or as paramagnetic shift probes producing narrow lines. Divalent manganese is a typical relaxation probe with an estimated electronic relaxation time (ts = T1c = Tc) of 10−8–10−9 s, while nickel, which has a shorter ts in the range 10−10–10−12 s, is a typical chemical shift probe. Cobalt(II) in a low-spin coordination environment has an estimated ts between that of Mn2+ and Ni2+ in kinetically labile metal complexes. At low metal to nucleotide ratios paramagnetic shift effects of Ni2+ are difficult to detect. In this case geometric information about metal binding sites is most effectively obtained by measuring proton spin-lattice relaxation times (T1). Paramagnetic relaxation arises in NMR spectroscopy when an unpaired electron spin interacts with a nuclear spin. The large magnetogyric ratio of the electron compared to that of the proton makes the dipolar coupling to the electron spin a very effective means of relaxation for the nuclear spin.25,26 Scalar interactions between the electron and nuclear spins have similar effects. In the simplest possible case, a ligand molecule exchanges between a paramagnetic environment (e.g. bound to Mn(II), S = 5/2) and a ‘free’ state, when the ligand is present in solution in vast excess to the paramagnetic centre (e.g. 102–104). The effect of paramagnetic metal ions located at specific binding sites on DNA is observed as differential linebroadening of proton signals close to the binding site. Often, in 1D spectra of oligonucleotide molecules containing ten base pairs or more, key proton resonances may be severely overlapped, preventing an accurate assessment of the influence of

10

Sequence-Selective Binding to DNA

the added metal ions. In these cases, 2D NOESY experiments may be used to obtain sufficient resolution. For diamagnetic metal ions (no unpaired spin) the formation of a chemical bond is usually found to cause changes in the chemical shifts of proton resonances of hydrogen atoms in the proximity of the metal binding site. However, the coordination of Hg2+ ions to single nucleobases induces only rather insignificant 1H shift changes, as shown for thymidine and guanosine.27 This could be explained by a down-field chemical shift change, induced by metal binding, being cancelled by an up-field chemical shift caused by changes in ring current effects due to altered nucleobase stacking.28 The heteronuclei 13C, 15N and 31P may experience large shifteffects when a metal ion binds to nucleobase or phosphate groups. 1.3.2 Model Systems One of the first oligodeoxynucleotides studied by X-ray crystallography29 and subsequently by NMR spectroscopy30 was the famous Dickerson–Drew sequence, d(C1G2C3G4A5A6T7T8C9G10C11G12) (seq. I). The 1D and 2D NMR spectra used as bases for the three-dimensional NMR structure showed an unexpectedly strong line-broadening of the G4-H8 resonance. As a consequence, the G4-H8 to C3-1′H cross-peak was barely detectable in the 2D NOESY spectra, and the corresponding interproton distance was scored as >4 Å by using the usual r6 dependency to calculate the proton–proton distance from the intensity of the cross-peaks.31 Structure determination based on protom–proton distance constraints showed a distinct bend of the helix axis at the G4 residue.30 However, at a later stage, when EDTA was added to the solution the G4-H8 resonance was found to exhibit normal line width.7 Since trace amounts of paramagnetic impurities are often found to be present in nucleotide preparations it is recommended to always run samples through a CHELEX column to remove such impurities before NMR metal ion titration experiments are performed. 1.3.3 Oligionucleotide–Transition Metal Adducts Mn2+ Ions The first metal ion titration NMR experiment on a DNA oligonucleotide was carried out by Frøystein and Sletten on the Dickerson–Drew duplex d(C1G2C3G4 A5A6T7T8C9G10C11G12) (seq. I) which contains the Eco-RI recognition sequence GAATTC.7 Since most of the base proton signals are well resolved in the 1D spectrum, the effect of adding aliquots of a solution of MnCl2 was easily observed. A plot of metal-induced paramagnetic line-broadening of G-H8 showed a clear preference for G4 compared to G2, G10 and G12 (Figure 1.5). At a Mn2+: duplex ratio of 1 : 104 substantial line-broadening was already detected for the G4-H8 signal, while insignificant effects were observed for the other base proton resonances in the duplex. Of particular importance was the observation that the central adenine A-H2 protons showed no broadening effect, indicating that Mn2+ ions were not located in

NMR Spectroscopic Studies

11

Figure 1.5 Mn2+ induced line-broadening of G-H8 resonances of the Dickerson–Drew duplex [d(5′-C1G2C3G4A5A6T7T8C9G10C11G12)]2. G4 (open squares), G10 (filled squares). The circles represent the overlapping G2 and G12 resonances7

the minor groove (see discussion below). The amount of broadening was consistent with inner sphere coordination to G-N7. The binding to guanine by 3d transition metals was not surprising, considering the differences in thermodynamic stability of the corresponding complexes of the nucleoside and nucleotide monomers.32 However, the sequence-selective metal binding pattern manifested through the apparent preference for G4 rather than the residues G2, G10 or G12, was highly unexpected. Further NMR spectroscopic evidence for apparent sequence-selective metal binding to DNA is presented for the duplex d(G1C2C3G4A5T6A7T8C9G10G11C12) (seq. II) containing the Eco-RV recognition sequence GATATC.8 The 1D spectra with and without MnCl2, shown in Figure 1.6, exhibit differential G-H8 line-broadening in the order: G10 > G4 >> G1 ∼ G11. It is evident that the exposed terminal residues both in sequence II and the previous sequence I, although offering favourable accessibility, are not the preferred binding sites. One may also notice that the A5-H2 and A7-H2 protons, residing in the minor groove, are completely unaffected. A systematic search for a selectivity pattern was initiated by designing three self-complementary sequences: [d(TATGGTACCATA)]2 (III), [d(TATGGATCCATA)]2 (IV) and [d(TATGGCCATA)]2 (V), where the central triplet is GGX (X = A, T or C).10 Mn2+-induced line-broadening for G-H8 versus r = [Mn2+]/[duplex] is plotted in Figure 1.7a–c. The line-broadening of G-H8 for the 5′-G is practically the same for all three sequences. However, the linebroadening of 3′-G-H8 is seen to depend on the adjacent residue X following a

12

Sequence-Selective Binding to DNA

Figure 1.6 400 MHz 1H NMR spectra, the aromatic region, of the dodecamer [d(GCCGATATCGGC)]2: (a) the spectrum of Mn2+-free solution, and (b) with added MnCl2 at a Mn2+/duplex ratio of approximately 10−3. The H8 and H6 proton resonances of purine and pyrimidine residues are labelled according to their sequential assignment8

distinct sequence-selective pattern: GA > GT >> GC. The adenine A-H8 resonances, plotted as references, are not influenced by the paramagnetic ions. Elmroth and coworkers have shown that metalation of single-stranded oligonucleotides of the type d(TnGT16−n) reach a maximum in the central part of the oligomer indicating lack of sequence-selective influence on the reaction rate.33,34 This result is in accordance with NMR and photo-cleavage studies on metalation of single-stranded DNA oligomers.18 Based on Monte Carlo descriptions of the oligoelectrolyte properties of double-stranded DNA oligomers it has been postulated that outer-sphere Coulombic interactions cause cations to be localized preferentially in the interior rather than the terminal part of the DNA oligomers.35 For inner-sphere metalation other factors may dominate, as demonstrated for the palindromic hexanucleotide d(G1G2C3G4C5C6) (seq. VI).36 Addition of MnCl2 induces selective line-broadening on the terminal G1-H8 with no significant effect on G2 and G5, as shown in the plot of line-broadening versus added MnCl2. In contrast, the terminal G1-H8 in duplex II (see above) shows almost no line-broadening. Labiuk et al.37 have reported X-ray determinations of Co2+, Ni2+ and Zn2+ complexes of d(G1G2C3G4C5C6) (seq. VI), where the metal ions are coordinated only to the terminal guanine G1-N7 position, with no metal ions binding to nonterminal guanine positions. The authors concluded that in the regular B-DNA conformation the internal binding sites are not accessible to Co2+, Ni2+ and Zn2+, and that consequently these metal ions bind exclusively to the terminal region of double-helical B-DNA, irrespective of base sequence. This is in contrast to our studies, which show a clear sequence-selective binding pattern for 3d metal ions with no special

NMR Spectroscopic Studies

13

Figure 1.7 Line-broadening versus Mn2+/Duplex ratio of the H8 resonances of the guanosines 5′-G4 (*) and 3′-G5 (ⵧ) with the terminal adenine H8 protons (䊊) for comparison. (a) d(TATGGTACCATA)2; (b) d(TATGGATCCATA)2; (c) d(TATGGCCATA)2. (J. Vinje, J.A. Parkinson, P.J. Sadler, T. Brown, E. Sletten, Sequence-selective metalation of double-helical oligodeoxyribonucleotides with PtII, MnII and ZnII ions. Chem. Eur. J., 2003, 9, 1620–1630. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Reproduced with permission.)

preference for terminal base residues. Based on the NMR results from several DNA sequences the d(GGCGCC) duplex is expected to bind preferentially to the 5′-G in the 5′-GG step and that the 5′-G in the 5′-GC step should have negligible affinity, in agreement with the X-ray result.37 Crystallographic data of a metal–DNA complex containing an internal GG step would probably show binding at this site. Table 1.1 is a compilation of available line-broadening data including both controlled titration experiments and the effect of paramagnetic impurities (reversed by addition of EDTA). One may notice that only Gs in the context G-purine exhibit maximum broadening. For sequence no. 6, which contains no G-purine step, the 5′-G-H8 resonance in the context of the 5′-GT step exhibits maximum broadening. A close inspection of Table 1.1 reveals that the line-broadening of 5′-G-H8 follows a consistent pattern, where the influence on the 5′-G-H8 is dependent on the residue

14

Sequence-Selective Binding to DNA Table 1.1 Mn2+ paramagnetic induced broadening of G-H8 resonances. The residues which show maximum broadening are marked in bold and underlined. Restriction enzyme recognition sequences are marked in italic Sequence 1. 5′-CGCGAATTCGCG 2. 5′-GCCGATATCGGC 3. 5′-GCCGTATACGGC 4. 5′-GCCAGATCTGGC 5. 5′-GAATTTAAATTC 6. 5′-CGCGTATACGCG 7. 5′-GCCGTGCACGGC 8. 5′-GCCGTTAACGGC 9. 5′-GCCTGATCAGGC 10. 5′-CCAAGCTTGG 11. 5′-GCCGAATTCGGC 12. 5′-ATGGGTACCCAT 13. 5′-TATGGTACCATA 14. 5′-TATGGATCCAAT 15. 5′-TATGGCCATA 16. 5′-GGCGCC

Eco-RI Eco-RV AccI BglII

HpaI BclI

on the 3′-side according to the simple rule : 5′-GG ≥ 5′-GA > 5′-GT >> 5′-GC. Apparently, the residue on the 5′-side is less important. At higher metal ion concentrations most 1H NMR resonances undergo varying degrees of broadening. It must be stressed, however, that only relative broadening effects for each sequence are taken into account. For sequences 12, 13 and 14 the GGX (X = A, T, G) triplet is placed in an almost identical environment. The effects of varying the X-residue are clearly shown in Figure 1.7. In sequence no. 12 containing a GGG triplet, both linebroadening and T1 data show the following order: G3 = G4 > G5 consistent for, not only Mn2+, but also for Ni2+ and Co2+ (see Ni2+ ions below). Fe2+ Ions In early investigations of Fe2+-DNA binding using NMR, the T1 spin–spin relaxation time technique indicated that both the base and the phosphate groups interact with Fe2+ ions.38 Later on, Bertoncini et al.39, by using X-ray absorption near edge structure spectroscopy (XANES), demonstrated that while Fe3+ tends to associate with oxygen ligands, Fe2+ prefers to form inner-sphere coordination with nitrogen ligands on DNA. Linn and coworkers have made extensive studies on sequence-specific DNA cleavage by Fe2+-mediated Fenton reactions.40–42 Nicking of duplex DNA by the iron-mediated Fenton reaction occurs preferentially at a limited number of nucleotide sequences. Most notable are a purine nucleotide followed by three or more G residues, [RGGG], and purine nucleotides flanking a TG combination, [TTGR].42 The preferred reaction sites are probably a consequence of sequenceselective localization of the ferrous ions. Using 1H NMR to characterize Fe2+ binding within the duplex CGAGTTAGGGTAGC/GCTACCCTAACTCG it was shown that Fe2+ binds preferentially at the GGG sequence, most strongly towards its 5′-

NMR Spectroscopic Studies

15

end.42 These studies are especially interesting because of the presence of RGGG in a large majority of telomere repeats.40 Recent studies have implicated that telomere shortening during human aging may be accelerated by oxidative stress.43,44 Co2+ Ions The Co2+ ion in a low-spin environment has an estimated electronic relaxation time (ts) between those of Mn2+ and Ni2+. Selective line-broadening was observed when CoCl2 was added to the duplex, d(A1T2G3G4G5T6A7C8C9C10A11T12) VII.9 At r = Co2+/ phosphate = 1.8 × 10−3 Co2+ was found to bind preferentially at the GGG sequence with almost equal affinity based on selective line-broadening; for G3 and G4 (35 Hz) and significantly less (12 Hz) for G5. NOESY spectra show that the intensities of cross-peaks were reduced by paramagnetic effects from the nearby bound Co2+ ion. The differential H8 broadening of G3, G4 and G5 observed in the 1D spectra is reflected as quenched NOE effects for the intra- and inter-proton cross-peaks involving G-H8 … G-H1′ connectivity. As a caveat we should mention that minute paramagnetic impurities in DNA samples used for NMR structure determination seriously affect the validity of the structural analysis since calculations of proton– proton distances are based on the magnitude of cross-peak intensities. Ni2+ Ions The electronic relaxation time (ts) for Ni2+ in the range 10−10 to 10−12 s is much shorter than for Mn2+. Consequently, spin-spin T1 relaxation time measurements will give more reliable information on metal binding sites than line-broadening (T2) data. Figure 1.8a shows that the pattern for Ni2+ binding to the Dickerson–Drew sequence deduced from T1 data is almost identical to that observed for Mn2+ binding based on line-broadening data.9 The T1 results of NiCl2 titration of the d(A1T2G3G4G5T6A7C8C9C10A11T12) duplex, which contains a triple GGG sequence are plotted in Figure 1.8b. The affinities for G3 and G4 are identical within experimental errors, while the affinity of G5 in the context GT is, as expected, significantly lower. The T1 relaxation pattern follows exactly the same trend as was shown for the Co2+ line-broadening measurements.9 Abrescia et al.45 reported an X-ray structure determination of the Ni2+ /DNA complex of the sequence d(CGTATATACG)2. The Ni2+ ions were associated with N7 atoms of all the guanines and with none of the N7 atoms of adenines. Subsequently the same group published a low resolution X-ray structure determination of the duplex d(CGTGTACACG), where they found Ni2+ ions associated only with the terminal Gs and the phosphate backbone.46 This last result is not in accordance with the sequence-selectivity observed in solution. However, the discrepancy may be due to crystal packing effects, which lead to cross-linking of guanines by Ni2+ ions. Cu2+ Ions One of the first detailed UV spectroscopic studies on the influence of metal ions on DNA stability was published by Eichhorn and Shin.3 While some metal ions tend

Sequence-Selective Binding to DNA

16 a

b 4

G4 H8

10

G3 H8 G4 H8

3

6 A5 H8

4

G2 H8 G12 H8 G10 H8

2 0 0

5

10

15 20 r*104

25

30

T1−1

T1−1

8

G5 H8

2 1 A11 H2

0

0

5

10

15 20 r*104

25

30

Figure 1.8 Plot of T1−1 versus r = [Ni(II)]/[Phosphate] at 312 K for G-H8 and A-H8 resonances of selected residues along the sequences: (a) [d(C1G2C3G4A5A6T7T8C9G10C11G12)]2; (b) [d(A1 T2G3G4G5T6A7C8C9C10A11T12)]2. (Reprinted from Inorg. Chim. Acta, 273, 1–2, E. Moldrheim, B. Andersen, N. A. Froystein, E. Sletten, Interaction of manganese (II), cobalt (II) and nickel (II) with DNA oligomers studied by 1H NMR spectroscopy, 6. Copyright 1998, with permission from Elsevier.)

to stabilize the double-helical structure, Cu2+ salts were shown to induce a dramatic lowering of the melting temperature (Figure 1.1). Recent NMR studies of Cu2+ and 2+ Cu( en)2 binding to the Dickerson–Drew duplex show the same selectivity pattern as for the other transition metals.47 The increase in line width at an r = [Cu2+]/ [Phosphate] of 2.5 × 10−3 was approximately 10 Hz, a corresponding increase in line width for [Cu(en)2]2+ was obtained at r = 15 × 10−3. Oikawa and Kawanishi have investigated the telomere shortening induced by H2O2 plus Cu2+ and found predominant DNA damage at the 5′-site of 5′-GGG-3′ in a 48-base fragment of the telomere duplex.48 In contrast, when single-stranded DNA was used, the damage induced by oxidative stress occurred at every guanine. The difference in site specificity of DNA damage between double-stranded DNA and single-stranded DNA could be explained in terms of the lower ionization potentials of stacked guanine base pairs in double-stranded DNA, on the basis of theoretical calculations. Zn2+ Ions UV spectroscopic titration studies of ctDNA by ZnCl2 indicate that Zn2+ ions bind both to phosphate groups and base nitrogen atoms.3 Marzilli and coworkers carried out Zn2+ titration of duplex d(ATG3G4G5TACCCAT) (VII) containing a GGG triplet, and found the chemical shift variation for the G-H8 in the order G4 > G3 >> G5.49 Zn2+ titration experiments analogous to those using Mn2+ (see above) have been carried out for sequences III, IV and V.10 In contrast to paramagnetic metal systems, where very low metal/DNA ratios were employed, excess salt was added to the duplex solutions until an upper limit of Zn2+-induced chemical shift changes

NMR Spectroscopic Studies

17

Figure 1.9 Chemical shift versus Zn2+/duplex ratio plots for the H8 resonances of 5′-G4 (*) and 3′-G5 (䊐), with the terminal adenine H8 protons (o) for comparison: (A) d(TATGGTACCATA)2; (B) d(TATGGATCCATA)2; (C) d(TATGGCCATA)2. (J. Vinje, J. A. Parkinson, P.J. Sadler, T. Brown, E. Sletten, Sequence-selective metalation of double-helical oligodeoxyribonucleotides with PtII, MnII and ZnII ions. Chem. Eur. J., 2003, 9, 1620–1630. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Reproduced with permission.)

was reached. This upper level was observed at r = [Zn2+]/[duplex] ≈ 5, which corresponds to a [Zn2+]/[G] value of ≈1.2. Plots of chemical shifts versus r exhibit the same trend as for the Mn2+ titration data (Figure 1.9). In a recent multinuclear NMR study on Zn2+ binding to oligonucleotides the variation in 1H, 15N and 31P chemical shifts was monitored as a function of added ZnCl2.36 Measurements for three different sequences were carried out: d(GGCGCC) (III); d(GGTACCGGTACC)2 (VIII); d(GGTATATATACCGGTATA) (IX). The chemical shift pattern for sequence III followed closely the line-broadening pattern determined for Mn2+ (see above). For sequence VIII large chemical 15N shift effects were observed for G7-N7 at natural abundance, a clear proof of direct metal binding to this site. 31P spectra also show large chemical shifts for the central G7 residue indicating that at excess ZnCl2 concentration the Zn2+ ions have direct contact with the G7 phosphate group and/or that a change in the phosphodiester backbone conformation has occurred.36

18

Sequence-Selective Binding to DNA

Zn2+ titration of IX was carried out to check if the expected accumulation of negative electrostatic potential in the interior of a long sequence33,34 (compared to shorter ones) would lead to a higher Zn2+ affinity for the internal GG step, compared with that observed for VIII.36 The results clearly showed that there is no enhanced selectivity for the internal Zn2+ binding for the long sequence. [Pt(dien)]+ Ions Numerous publications have dealt with the binding pattern of cisplatin anticancer drugs to DNA (see Chapters 5–9 in this book). One of the first studies on this topic involved enzymatic digestion of platinated DNA using the well known anticancer drug cisplatin (cis-[PtCl2(NH3)2]).50 HPLC analysis of the cleavage products showed that cisplatin forms chelates spanning G and/or A residues with the following percentages : Pt-GG (65%), Pt-AG (25%), Pt-GA (0%). Apparently, the rule of sequence-selectivity proposed for labile metal ion complexes based on 1H NMR and photochemical cleavage studies does not apply to nonlabile platinum complexes. However, the mechanism of chelate formation does not necessarily follow the sequence-selectivity of the initial formation of monofunctional adducts. The fact that the 5′-monoadducts are formed more rapidly and chelate more slowly than the 3′monoadducts might reflect the inherently greater reactivity of the 5′-G compared to the 3′-G. The common view is that the monoaqua species of cisplatin make the initial attack on DNA bases, although recent results have lead Chottard and coworkers to suggest that cisplatin may undergo double hydrolysis before reacting with DNA.51 In most reported studies on the kinetics of monofunctional platination reactions of single- and double-stranded oligonucleotides, both NMR spectroscopic and chromatographic methods have been used to determine kinetic parameters. The reactions involving bifunctional platinum complexes have usually been carried out with single-stranded oligonucleotides. In these reactions an initial monofunctional adduct is formed, which subsequently ring-closes to form a bifunctional chelate. The proposed selectivity rule is based on the intrinsic binding properties of duplex DNA. Sadler and coworkers showed that in a reaction between 15N-labelled [PtCl(dien)]+ and a single-stranded 14-mer d(ATACATGGTACATA), little kinetic preference for platination of either 5′G or 3′G sites was observed, while the single-stranded 8mer d(ATACATGG) showed a distinct preference for 5′-G platination.52 Chottard and coworkers, using hairpin-forming oligomers as duplex models, concluded that the selectivity for monofunctional attack by Pt2+ on 5′and 3′ G-residues is dependent on the ligand in the trans position (e.g. Cl−, H2O, OH−, NH3).51 In a later study they proposed a model for sequence-selective binding of cisplatin to DNA duplexes, involving a combination of molecular potentials and N7 accessibility.53 In a study on selectivity of adduct formation between 15N -cisplatin and 14-mer duplexes containing central AGT and GAT residues, respectively, Hambley and coworkers concluded that ‘the purine base on the 3′side of the pair exerts substantially greater influence on the rate of binding at the 5′-base than does the 5′-base on the rate of binding at the 3′-base’.54 However, monofunctional binding of cisplatin to the TGAT

NMR Spectroscopic Studies

19

Scheme 1.2

sequence was found to be approximately an order of magnitude slower than binding to TAGT sequences, a result which does not agree with the proposed selectivity rule for labile metal ion complexes. Vinje et al.10 have investigated the monofunctional reaction kinetics of [PtCl(dien)]+ (dien = diethylenetriamine), with the oligodeoxyribonucleotides, III, IV and V, used for Mn2+ and Zn2+ titration (see above). The reaction mixtures were separated by HPLC and the chromatographic profiles showed a clear difference in the amount of 3′ versus 5′ monoplatinated species between the three duplexes (Figure 1.10). The reaction pathway of platination of seq. III is shown in Scheme 1.2. Reaction rates for Pt(dien) were determined based on 2D [1H, 15N] HSQC/ HMQC NMR spectroscopy using 15N-labelled Pt(dien). The time courses for reaction rates of each of the three duplexes are shown in Figure 1.11. Comparison of the three different duplexes containing the central sections GGX (X = A, T, C), respectively,10 shows that the selectivity for covalent platination matches that for adducts with labile metal ions Mn2+ and Zn2+. The reaction rate for platination is faster for the 5′-G than for the 3′-G. The reactivity of the 3′-G depends on the adjacent residue X in the following order: X = A > T >> C. For GGA the reaction rate is 1.2 times faster for 5′-G than for 3′-G, for GGT the rate of 5′-G platination is about eight times faster than that of 3′-G, and for GGC there is no significant adduct formation at the 3′-G (Table 1.2). When the reaction mixture of the platinated species was aged over several weeks, the relative amounts of 5′ and 3′ HPLC fractions changed. This was also confirmed by comparison of the NMR spectra of the aged solutions with those recorded in the initial kinetic experiments. It was suggested that Pt-N7 bond cleavage and isomerization had taken place, similar to that observed for platinated single-

20

Sequence-Selective Binding to DNA

Figure 1.10 Chromatograms of the reaction products from the reaction between Pt(dien) and DNA: (a) d(TATGGTACCATA)2; (b) d(TATGGATCCATA)2; (c) d(TATGGCCATA)2. The HPLC fractions were collected ca 1 day after the reactions were started. The assignments of the fractions are based on analysis of 2D NOESY spectra. (J. Vinje, J.A. Parkinson, P.J. Sadler, T. Brown, E. Sletten, Sequence-selective metalation of double-helical oligodeoxyribonucleotides with PtII, MnII and ZnII ions. Chem. Eur. J., 2003, 9, 1620–1630. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Reproduced with permission.)

stranded DNA.52 Leng and coworkers have suggested a catalytic property of the DNA double helix to explain the rearrangement that is observed when a stable trans-platinated single strand is annealed with its complementary strand.54 The occurrence of Pt–N bond cleavage may influence the results of kinetic analysis that are based on HPLC techniques, in which aliquots of the reaction mixture are collected at several time points and quenched with large amount of potassium chloride.

NMR Spectroscopic Studies

21

Figure 1.11 Experimental concentrations (NMR data) and theoretically-fitted curves for the reactions between [Pt(dien)]+ and DNA: (a) d(TATGGTACCATA)2; (b) d(TATGGATCCATA)2; (c) d(TATGGCCATA)2. Symbols: (ⵧ) Pt(dien); (o) Pt-G5′and (∆) Pt-G3′. (J. Vinje, J.A. Parkinson, P.J. Sadler, T. Brown, E. Sletten, Sequence-selective metalation of double-helical oligodeoxyribonucleotides with PtII, MnII and ZnII ions. Chem. Eur. J., 2003, 9, 1620–1630. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Reproduced with permission.)

Table 1.2 Rate constants (standard deviations in parentheses) for reactions of [PtCl(dien)]+ with DNA oligonucleotides. pH 5.6, 0.1 M NaClO410 Sequence

k5′ (M−1 s−1)

k3′ (M−1 s−1)

-GGTA-(a) -GGAT-(a) -GGCC-(b)

4.3 (6) 4.0 (6) 7.6 (6)

0.5 (1) 3.4 (5) (c)

Temperature: (a) 288 K; (b) 298 K; (c) Negligible 3′-G platination.

22

Sequence-Selective Binding to DNA

1.4 Summary of Theoretical and Experimental Evidence for Sequence–Selective Binding to DNA Paramagnetic and diamagnetic ions are shown to form both labile and nonlabile adducts with DNA duplexes, and in a series of G-containing duplexes a certain selectivity pattern emerges where the metal prefers the 5′-G in the following order: 5′-GG > 5′-GA > 5′-GT >> 5′-GC. The adjacent residue (Y) on the 5′-side (5′-YGG) is found to exert a negligible influence on the selectivity. For the monofunctional binding of [PtCl(dien)]+ to double helical DNA the variation in reaction rates follows qualitatively the same selectivity pattern as for the labile metal ions. Experimentally observed relative rates of G-oxidation are found to match well with the NMR results. Nonselective equal G cleavage is observed for single-stranded DNA, in line with the NMR experimental data. Ab initio molecular orbital calculations of stacked DNA bases with B-form geometry clearly indicated that the highest occupied molecular orbital (HOMO) localization on the 5′-G is highly sequencedependent. The degree of p–p interaction between the stacked bases influences the HOMO energies. However, it should be emphasized that for metal complexes the stereochemistry of the ligands also plays a significant role in determining the most favourable binding site.

1.5 Sequence-Specific Groove Binding Among the large number of contributions to the field of cation–groove interaction, probably a majority is based on X-ray crystallography. Sequence-specific nucleic acid conformation and dynamics are directly influenced by metal ions. DNA conformational heterogeneity has been explained by an electrostatic model where the local position and transient fluctuation of ions act through asymmetric neutralization of phosphate charges. Evidence from NMR spectroscopy, X-ray crystallography and molecular dynamics simulations has revealed that B-form duplexes interact in a sequence-specific manner with fully hydrated mono and divalent cations.55 1.5.1 Groove Geometry A noteworthy feature of B-DNA is the presence of two kinds of grooves, called the major groove (12 Å wide) and the minor groove (6 Å wide) (see figures in biochemistry text books). They arise because the glycosidic bonds of a base pair are not diametrically opposite each other. The minor groove contains the pyrimidine O2 and the purine N3 of the base pair, and the major groove is on the opposite side of the pair. In the minor groove, N3 and O2 can serve as hydrogen bond acceptors, and the amino group attached to guanine can be a hydrogen donor. In the major groove N7 is a potential hydrogen bond acceptor, as are O4 of thymine and O6 of guanine. The amino groups attached to adenine and cytosine, respectively, can serve as hydrogen donors.

Sequence-Specific Groove Binding 23

The width of the grooves is not constant, but is found to be sequencedependent. Alternative models have been presented to provide explanations for this sequence-dependent variation. NMR spectroscopic analyses of a series of DNA duplexes have shown that alternating (AT)n sequences are characterized by a rather wide minor groove.56,57 From a number of experimental and theoretical studies, it has been concluded that on average the minor groove of DNA A-tracts (several consecutive adenine residues) is significantly narrower than the minor groove of G-tracts.58–60 1.5.2 Monovalent Cations In the report of the first X-ray structure of a double-helical DNA duplex, [d(CGCGAATTCGCG)]2, published by Dickerson and Drew,58 it was proposed that a ‘spine of hydration’, composed of localized, geometrically arranged water molecules, in the minor groove, is an important structural component of A-tract DNA. However, this result is not fully conclusive since Na+ and H2O, being isoelectronic, are not easily distinguishable by X-ray diffraction. Egli and coworkers repeated the structural analyses of the same duplex using X-ray diffraction data to near-atomic resolution with crystals grown in the presence of Rb+ cacodylate.61 They found that a single Rb+ ion, with partial occupancy, was localized at the central ApT step at the bottom of the AATT minor groove (Figure 1.12). The ion replaces the water molecules that link the keto oxygen of thymines from opposite strands. The authors suggest that minor groove ion coordination appears to be an isolated event, highly sequence-dependent and unlikely to significantly affect the particular geometry of the A-tract in the Dickerson–Drew dodecamer. Further studies by Williams and coworkers on the same duplex using Tl+ to mimic K+ showed that none of the observed Tl+ sites surrounding the duplex were fully occupied.62 The

Figure 1.12 Coordination of Rb+ at the central ApT step in the Dickerson–Drew Duplex.61 (Reprinted from Chem. Biol., 9, 3, M. Egli, DNA-cation interactions: Quo vadis?, 10. Copyright 2002, with permission from Elsevier.)

24

Sequence-Selective Binding to DNA

most highly occupied sites (20–35%) were located within the G-tract major groove while the occupancy in the minor groove were estimated to be around 10%. The situation concerning penetration of Na+ ions into the spine of hydration is still an open question among X-ray crystallographers. The existence of just a single alkali metal ion coordination site in the Rb+-form crystal structure argues against a view that ions can invade the minor groove hydration spine along the entire length, or the existence of a mixed water-ion spine of hydration. In a recent high-resolution (1.1 Å) structure determination of the Dickerson–Drew duplex no experimental evidence for the presence of Na+ ions in the minor groove was found.63–65 A series of 23Na NMR quadrupolar relaxation studies have been carried out on B-DNA.66,67 A general conclusion from these studies is that monovalent counterion binding to DNA is loose and delocalized, without any dehydration or sequencespecific features. In an optimally designed test for sequence-specific Na+ binding in the minor groove Denisov and Halle have used a magnetic relaxation dispersion (MRD) technique, where the 23Na relaxation rate is measured over nearly two decades of resonance frequency.67 Comparison of Na+ MRD data from three dodecamers with different nucleotide sequences: CGCGAATTCGCG (abbreviated A2T2), CGAAAATTTTCG (A4T4) and CGCTCTAGAGCG (TA) showed that the most tightly bound Na+ ions reside in the minor groove. However, the occupancies are quite low corresponding to a binding constant KNa of 0.03 M−1 for TA and 0.1 M−1 for the other two dodecamers, and imply that Na+ binding in the minor groove is a rare event and is not likely to be detected by X-ray diffraction. These results are not necessarily inconsistent with higher occupancy at the cryogenic temperature (120–160 K) used in recent crystallographic studies. Even a modest binding enthalpy of 5 kJ mol−1 could increase the binding constant from 0.1 M−1 at 277 K to 1.7 M−1 at 120 K, which is sufficient to give 50% occupancy in a single binding site, as found for Rb+ at the ApT step in A2T2.61 The authors conclude that groove bound Na+ ions, with an occupancy of only a few percent at room temperature, are not likely to contribute importantly to the ensemble of DNA structures under physiological conditions.67

1.5.3 Divalent Cations At physiological concentrations the binding of divalent cations to DNA is both cation dependent and sequence dependent. From a sequence standpoint, specificity is contributed to by both the local molecular nucleophilicity (see above) and the hydrogen bond environment. These H-bond interactions reflect the greater hydration properties of divalent cations over monovalent. Generally, it is difficult to assess how contributions of base sequence or cation type influence groove specificity because of crystal packing effects. Chiu and Dickerson have examined a database of 28 cation-bound B-DNA structures spanning ten different crystal packing environments and showed that there is a correlation between experimental conditions and the number of observed cations.68 Hence, the locations of cations can be compared safely only between structures having similar crystallizing conditions, data collection methods and resolutions. Despite these difficulties, the authors find a

Sequence-Specific Groove Binding 25

Figure 1.13 Coordination of Mg2+ at the CpG step near one end of the Dickerson–Drew Duplex.63 (Reprinted from Chem. Biol., 9, 3, M. Egli, DNA-cation interactions: Quo vadis?, 10. Copyright 2002, with permission from Elsevier.)

strong correlation between divalent cation binding and base sequence for the 28 structures examined. For four high-resolution (0.99 Å) structures, the minor groove affinity for Mg2+ is GG > AG > AC and for Ca2+ is GG > AT ∼ AC, with cations positioned in the centre of the groove. For the major groove, the order for Mg2+ is GG > AG ∼ GT and for Ca2+ is GG > AG.68 Treshko et al.63 have redetermined the crystal structure of the Dickerson–Drew duplex at 1.1 Å resolution. Three ordered Mg2+ ions are present in the asymmetric unit, two hexahydrates and one pentahydrate complex. One Mg2+ is located in the major groove, close to the end of the duplex (Figure 1.13). The ion contacts the N7 and O6 edges of residue G2 and G22 from opposite strands via coordinated water molecules. None of the contacts between Mg2+ ions and DNA atoms on the floor of the major groove involve inner-sphere coordination.65 Hud and Feigon have studied the localization of Mn2+ in A-tract DNA by 1H NMR spectroscopy, using a series of self-complementary dodecamer oligonucleotides that contain the sequence motifs AnTn and TnAn, where n = 2, 3 or 4 flanked by 5′-CG or 5′-GC base pairs.69 At an Mn2+ to duplex ratio of 10−3 most of the aromatic base protons are severely broadened, however, in contrast to the Mn2+ NMR data referred to above, the adenine H2 resonances exhibit the largest broadening. The authors conclude that Mn2+ is localized in the minor groove with the position and degree of localization being highly sequence-dependent. In addition, G2-H8 in the sequence CGT is broadened by 7 Hz/µM MnCl2 while G1-H8 in the sequence GCT is broadened less than 2 Hz, a finding that is in agreement with the observation that the affinity of 5′-GT >> 5′-GC (see above).

26

Sequence-Selective Binding to DNA

NMR spectroscopy70 and molecular dynamics (MD)71 studies have shown that AnTn and TnAn duplexes have unusual structures and dynamics. Anomalous broadening of the TpA adenine H2 resonance indicative of large amplitude base motion, has been observed for a series of nine unique four-nucleotide sequences.70 In AnTn sequences, the DNA assumes a unique structure characterized by a gradual and increasingly compressed minor groove, which reaches a minimum at the ApT step. In conclusion, this type of sequences does not exhibit a regular B-form conformation and thus minor groove cation binding may be unique to A-tract sequences. 1.5.4 Conclusion on Groove Binding The general trend is that ions bind in the minor groove of DNA A-tracts and in the major groove of DNA G-tracts. NMR spectroscopic studies indicate that sequencespecific binding of alkali and alkali earth ions appears to have only minor influence on the heterogeneity of DNA structures.67. The relative lack of hydration of alkali cations makes their interactions with base atoms largely electrostatic and relatively nonspecific. Divalent alkali earth cations tend to be fully hydrated and their interactions with duplex DNA are more sequence-specific through formation of hydrogen bonds to base atoms.55,67

Abbreviations MEP HOMO LUMO ODN MD EDTA XANES NOESY MRD

Molecular electrostatic potential Highest occupied molecular orbital Lowest occupied molecular orbital Oxidation oligodeoxynucleotides Molecular dynamics Ethylene diamine tetraacetic acid X-ray absorption near edge structure Nuclear overhauser effect spectroscopy Magnetic relaxation dispersion

References 1. Flessel, C.P.; Furst, A.; Radding, S.B.; A comparison of carcinogenic metals. In: Metal Ions in Biological Systems. Vol. 10, H. Sigel, Ed., Marcel Dekker, New York, 1980. 2. Frieden, E.; Alles, J.; Subtle interactions of cupric ions with nucleic acid components; J. Biol. Chem., 1958, 230, 797–804. 3. Eichhorn, G.L.; Shin, Y.A.; Interaction of metal ions with polynucleotides and related compounds. XII. The relative effect of various metal ions on DNA helicity; J. Am. Chem. Soc., 1968, 90, 7323–7328. 4. Sletten, E.; Lie, B.; Copper complex of guanosine-5′-monophosphate; Acta. Cryst., 1976, B32, 3301–3304.

References

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5. Yamane, T.; Davidson, N.; On the complexing of deoxyribonucleic acid (DNA) by mercuric ion; J. Am. Chem. Soc., 1961, 83, 2599–2607. 6. Frøystein, N.Å.; Sletten, E.; Interaction of mercury(II) with the DNA dodecamer CGCGAATTCGCG. A 1H and 15N NMR study; J. Am. Chem. Soc., 1994, 116, 3240–3250. 7. Frøystein, N.Å.; Sletten, E.; The binding of manganese(II) and zinc(II) to the synthetic oligonucleotide [d(CGCGAATTCGCG)]2; Acta Chem. Scand., 1991, 45, 219–225. 8. Frøystein, N.Å.; Davis, J.T.; Reid, B.R.; Sletten, E.; Sequence-selective metal ion binding to DNA oligonucleotides; Acta Chem. Scand., 1993, 47, 649–657. 9. Moldrheim, E.; Andersen, B.; Frøystein, N.Å.; Sletten, E.; Interaction of manganese(II), cobalt(II) and nickel(II) with DNA oligomers studied by 1H NMR spectroscopy; Inorg. Chim. Acta., 1998, 273, 41–46. 10. Vinje, J.; Parkinson, J.A.; Sadler, P.J.; Brown, T.; Sletten, E.; Sequence-selective metalation of double-helical oligodeoxyribonucleotides with PtII, MnII and ZnII ions; Chem. Eur. J., 2003, 9, 1620–1630. 11. (a) Laing, L.G.; Gluic, T.C.; Draper, D.E.; Stabilization of RNA structure by Mg ions. Specific and nonspecific effects; J. Mol. Biol., 1994, 237, 577–587. (b) Misra, V.; Draper, D.E.; A thermodynamic framework for Mg2+ binding to RNA; PNAS, 2001, 98, 12456– 12461. (c) Manning, G.S.; The molecular theory of polyelectrolyte solutions with applications to the electrostatic properties of polynucleotides; Q. Rev. Biophys., 1978, 11, 179–246. (d) Strehlow, H.; Rapid Reactions in Solution, VCH, Weinheim, 1992. 12. Black, C.B.; Cowan, J.A.; Quantitative evaluation of electrostatic and hydrogen-bonding contributions to metal cofactor binding to nucleic acids; J. Am. Chem. Soc., 1994, 116, 1174–1178. 13. Kohn, K.W.; Hartley, J.A.; Mattes, W.B.; Mechanisms of DNA sequence selective alkylation of guanine-N7 positions by nitrogen mustards; Nucleic Acids Res., 1987, 15, 10531–10549. 14. Monjardet, V.; Elizondo-Riojas, M.-A.; Chottard, J.-C.; Kozelka, J.; A combined effect of molecular electrostatic potential and N7 accessibility explains sequence-dependent binding of cis-[Pt(NH3)2(H2O)2]2+ to DNA duplexes; Angew. Chem. Int. Ed., 2002, 41, 2998–3001. 15. Fukui, K.; Recognition of stereochemical paths by orbital interaction; Acc. Chem. Res., 1971, 4, 57–64. 16. Hutter, M.; Clark, T.; On the enhanced stability of guanine-cytosine base-pair radical cation; J. Am. Chem. Soc., 1996, 118, 7574–7577. 17. Sugiyama, H.; Saito, I.; Theoretical studies of GG-specific photocelavage of DNA via electron transfer: significant lowering of ionization potential and 5′-localization of HOMO of stacked GG bases in B-form DNA; J. Am. Chem. Soc., 1996, 118, 7063–7068. 18. Saito, I.; Takayama, M.; Sugiyama, H.; Nakatani, K; Tsuchida, I; Yamamoto, M.; Photoinduced DNA cleavage via electron transfer: demonstration that guanine residues located 5′ to guanine are the most electron-donating sites; J. Am. Chem. Soc., 1995, 117, 6406–6407. 19. Nakamura, T.; Nakatani, K.; Saito, I.; DNA HOMO as a new landmark for nucleic acid properties. Ab initio calculations and experimental mapping; Nucleic Acids Res., 1999, 42, 119–120. 20. Saito, I.; Nakamura, T.; Nakatani, K.; Mapping of highest occupied molecular orbitals of duplex by cobalt-mediated guanine oxidation; J. Am. Chem. Soc., 2000, 122, 3001–3006. 21. Senthikumar, K.; Grozema, F.C.; Guerra, C.; Bickelhaupt, F.M.; Siebbeles, L.D.A.; Mapping the sites for selective oxidation of guanines in DNA; J. Am. Chem. Soc.; 2003, 125, 13658–13659. 22. Saito, I.; Nakamura, T.; Nakatani, K.; Yoshioka, Y.; Yamaguchi, K.; Sugiyama, H.; Mapping of the hot spots for DNA damage by one-electron oxidation: efficacy of GG doublets and GGG triplets as a trap in long-range hole migration; J. Am. Chem. Soc., 1998, 120, 12686–12687.

28

Sequence-Selective Binding to DNA

23. Cleveland, C.L.; Barnett, R.N.; Bongiorno, A.; Joseph, J.; Liu, C.; Schuster, G.B.; Landman, U.; Steric effects on water accessibility control sequence-selectivity of radical cation reactions in DNA; J. Am. Chem. Soc., 2007, 129, 8408–8409. 24. Behling, R.W.; Kearns, D.R.; Proton two-dimensional nuclear Overhauser effect and relaxation studies of poly(dA).cntdot.poly(dT); Biochemistry, 1986, 25, 3335–3346. 25. Kowalewski, J.; Nordenskiöld, L.; Benetis, N.; Westlund, P-O.; Theory of nuclear spin relaxation in paramagnetic systems in solution; Prog. NMR Spectrosc., 1985, 17, 141–185. 26. Navon, G.; Valensin, G.; Nuclear relaxation times as a source of structural information. In: Metal Ions in Biological Systems, Vol. 21. H. Sigel, Ed. Marcel Dekker, New York, 1987. 27. Buncel, E.; Boone, C.; Joly, H.; Metal ion-biomolecule interactions. Part 13. NMR evidence for the formation of the mixed ligand thymidine-mercury-guanosine complex. A model for a putative Hg(Il) interstrand cross-linking structure of DNA; Inorg. Chim. Acta, 1986, 125, 167–172. 28. Young, P.R.; Nandi, U.S.; Kallebach, N.R.; Binding of mercury(II) to poly(dA-dT) studied by proton NMR; Biochemistry, 1982, 21, 62–66. 29. Drew, H.; Dickerson, R.E.; Structure of a B-DNA dodecamer. III. Geometry of hydration; J. Mol. Biol., 1981, 151, 535–556. 30. Nerdal, W.; Hare, D.R.; Reid, B.R.; Solution structure of the Eco-RI DNA sequence: refinement of NMR-derived distance geometry structures by NOESY spectrum backcalculation; Biochemistry, 1989, 28, 10008–10021. 31. Hare, D.R.; Reid, B.R.; Three-dimensional structure of a DNA hairpin in solution: twodimensional NMR studies and distance geometry calculations on d(CGCGTTTTCGCG); Biochemistry, 1986, 25, 5341–5350. 32. Eichhorn, G.L.; Complexes of nucleosides and nucleotides. In: Eichhorn, G. L., Ed. Inorganic Biochemistry, Elsevier, Amsterdam, 1973, Vol. 2, Chap. 33, 1192–1209. 33. Snygg, Å.S.; Ericson, A.; Elmroth, S.K.C.; Nonuniform rate of platination of guanine-N7 located in short DNA oligomers; Chem. Commun., 2001, 1190–1191. 34. Snygg, Å.S.; Brindell, M.; Stochel, G.; Elmroth, S.K.C.; A combination of access to preassociation sites and local accumulation tendency in the direct vicinity of G-N7 controls the rate of platination of single-stranded DNA; Dalton Trans., 2005, 1212–1227. 35. Olmsted, M.C.; Anderson, C.F.; Record, M.T.; Monte Carlo description of oligoelectrolyte properties of DNA oligomers: range of the end effect and the approach of molecular and thermodynamic properties to the polyelectrolyte limits; Proc. Natl. Acad. Sci., 1989, 86, 7766–7770. 36. Vinje, J.; Sletten, E.; Internal versus terminal metalation of double-helical oligodeoxyribonucleotides; Chem. Eur. J., 2006, 12, 676–688. 37. Labiuk, S.L.; Delbaere, L.T.J.; Lee, J.S.; Cobalt(II), nickel(II) and zinc(II) do not bind to intra-helical N(7) guanine positions in the B-form crystal structure of d(GGCGCC); J. Biol. Inorg. Chem., 2003, 8, 715–720. 38. Eisinger, J.; Schulman, R.G.; Szymanski, B.M.; Transition metal binding in DNA solutions; J. Chem. Phys., 1962, 36, 1721–1729. 39. Bertoncini, C.; Meneghini, R.; Cruz, D.Z.; Alves, M.C.M.; Tolentino, H.; Studies of Fe(II) and Fe(III)-DNA complexes by XANES spectroscopy. J. Synchtron Rad., 1999, 6, 417–418. 40. Henle, E.S.; Han, Z.; Tang, N.; Rai, P.; Luo, Y.; Linn, S.; Sequence-specific DNA cleavage by Fe2+-mediated Fenton reactions has possible biological implications; J. Biol. Chem., 1999, 274, 962–971. 41. Rai, P.; Cole, T.D.; Wemmer, D.E.; Linn, S.; Localization of Fe2+ At an RTGR sequence within a DNA duplex explains preferential cleavage by Fe2+ and H2O2; J. Mol. Biol., 2001, 312, 1089–1101. 42. Rai, P.; Wemmer, D.E.; Linn, S.; Preferential binding and structural distortion by Fe2+ at RGGG-containing DNA sequences correlates with enhanced oxidative cleavage at such sequence; Nucl. Acid. Res., 2005, 33, 497–510.

References

29

43. Saretzki, G.; von Zglinicki, T.; Replicative aging, telomeres and oxidative stress; Ann. N.Y. Acad. Sci., 2002, 959, 24–29. 44. Kawanishi, S.; Hiraku, Y.; Oikawa, S.; Mechanism of guanine-specific DNA damage by oxidative stress and its role in carcinogenesis and aging; Mutant. Res., 2001, 488, 65–76. 45. Abrescia, N.G.A.; Malinina, L.; Fernandez, L.G.; Huybh-Dinh, T.; Neidle, S.; Subirana, J.A.; Structure of the oligonucleotide d(CGTATAATACG) as a site-specific complex with nickel ions; Nucl. Acid. Res., 1999, 27, 1593–1599. 46. Abrescia, N.G.A.; Huybh-Dinh, T.; Subirana, J.A.; Nickel-guanine interactions in DNA: crystal structure of nickel-d[CGTGTACACG]2; J. Biol. Inorg. Chem., 2002, 7, 195–199. 47. Emwas, A.H.M.; Sletten, E.; NMR studies of interaction between Cu2+ and Cu(en)2 with the Dickerson–Drew duplex [d(CGCGAATTCGCG)]2. Manuscript (to be published). 48. Oikawa, S.; Kawanishi, S.; Site-specific DNA damage at GGG sequences by oxidative stress may accelerate telomere shortening; FEBS Lett., 1999, 453, 365–368. 49. Jia, X.; Zon, G.; Marzilli, L.G.; Multinuclear NMR investigation of zinc(2+) binding to a dodecamer oligodeoxyribonucleotide: insights from carbon-13 NMR spectroscopy; Inorg. Chem., 1991, 30, 228–239. 50. Fichtinger-Schepman, A.M.; Van der Veer, J.L.; den Hartog, J.H.J.; Reedijk, J.; Adducts of the antitumor drug cis-diamminedichloroplatinum(II) with DNA: formation, identification, and quantitation; Biochemistry, 1985, 24, 707–713. 51. Legendre, F.; Monjardet-Bas, V.; Kozelka, J.; Chottard, J.C.; A complete kinetic study of GG versus AG platination suggests that the doubly aquated derivatives of cisplatin are the actual DNA binding species; Chem. Eur. J., 2000, 6, 2002–2010. 52. Murdoch, P.D.; Guo, Z.J.; Parkinson, J.A.; Sadler, P.J.; Kinetics of formation and stability of {Pt(dien)}2+ complexes with octamer and 14-mer DNA oligonucleotides containing a GG sequence; J. Biol. Inorg. Chem., 1999, 4, 32–38. 53. Davis, M.S.; Berners-Price, S.J.; Hambley, T.W.; Rates of platination of AG and GA containing double-stranded oligonucleotides: insights into why cisplatin binds to GG and AG but not GA sequences in DNA; J. Am. Chem. Soc., 1998, 120, 11380–11390. 54. Dalbies, R.; Payet, D.; Leng, M.; DNA double helix promotes a linkage isomerisation reaction in trans-diamminedichloroplatinum(II)-modified DNA; Proc. Natl. Acad. Sci., 1994, 91, 8147–8151. 55. Hud, N.V.; Polak, M.; DNA-cation interactions: the major and minor grooves are flexible ionophores; Curr. Opin. Struct. Biol., 2001, 11, 293–301. 56. Chuprina, V.P.; Lipanov, A.A.; Fedoroff, O.Y.; Kim, S-G.; Kintanar, A.; Reid, B.R.; Sequence effects on local DNA topology; Proc. Natl. Acad. Sci., 1991, 88, 9087–9091. 57. Chuprina, V.P.; Sletten, E.; Fedoroff, O.Y.; Investigation of solution structure of d(GAATTTAATTC)2 by 1H NMR, molecular dynamics, mechanics, refinement by backcalculation of the NOESY spectrum and analysis of this structure using X-ray data; J. Biomol. Struct. Dyn., 1993, 10, 693–707. 58. Dickerson, R.F.; Drew, H.R.; Structure of a B-DNA dodecamer.II. Influence of base sequence on helix structure. J. Mol. Biol., 1981, 149, 761–786. 59. Burkhoff, A.M.; Tullius, T.D.; The unusual conformation adopted by the adenine tracts in kinoplast DNA; Cell, 1987, 48, 935–943. 60. Alexeev, D.G.; Lipanov, A.A.; Skuratovskii, I.Y.; Poly(dA)-poly(dT) is a B-type double helix with distinctively narrow minor groove; Nature, 1987, 325, 821–823. 61. Tereshko, V.; Minasov, G.; Egli, M.; A ‘hydrated’ spIne in a B-DNA minor groove; J. Am. Chem. Soc., 1999, 121, 3590–3595. 62. Howerton, S.B.; Sines, C.C.; VanDerveer, D.; Williams, L.D.; Locating monovalent cations in the groove of B-DNA; Biochemistry, 2001, 40, 10023–10031. 63. Tereshko, V.; Minasov, G.; Egli, M.; The Dickerson–Drew B-DNA dodecamer revisited at atomic resolution; J. Am. Chem. Soc., 1999, 121, 470–471. 64. Tereshko, V.; Minasov, G.; Egli, M.; Atomic-resolution crystal structure of B-DNA reveal specific influence of divalent metal ions on conformation and packing; J. Mol. Biol., 1999, 291, 83–99. 65. Egli, M.; DNA-cation interactions: Quo vadis? Chem. Biol., 2002, 9, 277–286.

30

Sequence-Selective Binding to DNA

66. Halle, B.; Denisov, V.P.; Water and monovalent ions in the minor groove of B-DNA oligonucleotides as seen by NMR; Biopolymers, 1998, 48, 210–233. 67. Denisov, V.P.; Halle, B.; Sequence-specific binding of counterions to B-DNA; Proc. Natl. Acad. Sci., 2000, 97, 629–633. 68. Chiu, T.K.; Dickerson, R.E.; 1 Å crystal structures of B-DNA reveal sequence-specific binding and groove-specific bending of DNA by magnesium and calcium; J. Mol. Biol., 2000, 301, 915–945. 69. Hud, N.V.; Feigon, J.; Characterization of divalent cation localized in the minor groove of AnTn and TnAn DNA sequence elements by 1H NMR spectroscopy and managnese(II); Biochemistry, 2002, 41, 9900–9910. 70. McAteer, K.; Ellis, P.D.; Kennedy, M.; The effects of sequence context on base dynamics at TpA steps in DNA studied by NMR; Nucleic Acids Res., 1995, 23, 3962–3966. 71. Leporc, S.; Mauffret, O.; Tevanian, G.; Lescot, E.; Monnot, M.; Fermandjian, S.; An NMR and molecular modelling analysis of d(CTACTGCTTTAG).d(CTAAAGCAGTAG) reveals that the particular behaviour of TpA steps is related to edge-to-edge contacts of their base-pairs in the major groove; Nucleic Acids Res., 1999, 27, 4759–4767.

2 Thermodynamic Models of Metal Ion–DNA Interactions Vasil Bregadze, Eteri Gelagutashvili and Ketevan Tsakadze

2.1 Introduction Recently the catalytic properties of DNA became of special interest to scientists. In particular, attention has been drawn to photo-processes1–7 and processes connected with charge8–15 and energy16–18 transfers. On one hand, transition- (G ↔ A, C ↔ T) and transversion- (A ↔ T, A ↔ C, G ↔ C, G ↔ T) type gene or point mutations, in rare cases, may lead to improvement of one or another characteristic of the organism. They are essential steps for artificial or natural selection in biology. On the other hand, more commonly, these mutations cause transformation of the cells, or their death19–20. This fact finds its application in medicine, specifically in chemotherapy and photo-chemotherapy. The mutations that lead to death of the cells are initiated by anticancer drugs, which often contain metals.4–7,21–25 The structure of the DNA double helix offers carte blanche for transition-type mutations via defects, such as wrong Watson–Crick pairs, where self-congruent cooperative keto–enol and amino–imine tautomeric transitions, named doubleproton transfer (DPT), take place.17 This is a property of cyclic structures in polar molecules. The nucleotide pairs in DNA naturally present such cyclic structures between complementary bases. The mechanism of DPT in a DNA duplex was first considered by Löwdin as early as in 1963.26,27 It is assumed that approximately one Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

32

Thermodynamic Models

− Table 2.1 Reaction rate constants of eaq with H3O+ and transition metal ions. (Values of Ke/aq 9 −1 −1 are given in 10 m s .)

Mn+

H+

Mn2+

Co2+

Ni2+

Ni(en)2+

Ni(en3)2+

Cu2+

Cu+

Zn2+

Ag+

Ke/aq

24

0.038

12

22

22.4

0.02

30

–

1.5

36

DPT takes place spontaneously in one out of 104 bp. Transition metal ions are known to interact with DNA (depending on anion charge, G-C composition of DNA, ionic strength etc.28), the effect of which is to increase the number of mispaired Watson– Crick base pairs. The interaction of H3O+ and transition metal ions with DNA depends on both the electronic structure of the ions and the structural, polyelectrolytic and dynamic characteristics of DNA in water solution. − Table 2.1 presents the reaction rate constants of hydrated electrons (eaq ) with 29,30 different types of ions. The reactivities of the ions presented in the table actually reflect the affinities of the hydrated electron with these ions in water solutions31 and, as will be shown in Section 2.3.3, they correlate well with the results of UV spectra of DNA–metal ion complexes. Native double helix DNA is a heavily charged polyanion (2e− charge per base pair), which in solution exists in the form of an ordered closely packed structure, as proved by the strong hyperchroism displayed upon melting of the double helix. Quantum chemical evaluations show an identical sequence of arrangement of electron-donor atoms for unscreened B-DNA and for that screened by Na+ ions:32–33 G-N7; G-N3; A-N3; T-O2; G-O6; A-N7 The NH2 group of adenine and guanine prevents interaction of metal ions with A-N7 and G-N3 due to steric crowding. In addition, G-N3, A-N3 and T-O2 are located in the minor groove, the width and the depth of which are 5.7 Å and 7.5 Å, respectively, in the B-form. For steric reasons all these atoms are unreachable for a hydrated transition metal ion, which has a diameter of approximately 7.5 Å. However, the omnipresent H3O+ ion can interact with electron-donor groups from both of the grooves. Interaction of ions with DNA also depends on ionic strength, GC composition, the order of the nucleotides in the helix and the conformational parameters of DNA (the angle of the coil twist, the propeller angle, the distance between adjacent phosphorus atoms of the same chain etc.). Nevertheless, one may say with a certain assurance that dynamic characteristics of DNA are the most significant factors governing metal ion–DNA interactions. First of all, this is important for modelling e.g. B→Z transitions, mechanisms of transcription, replication, carcinogenesis and mutagenesis. In this respect it is important to describe the characteristics of inner mobility in DNA, e.g. various small-amplitude oscillating movements of atoms, ′ inside the components of DNA (10−14–10−13 s); limited movements of about 0.1 Å phosphates, sugars and nucleobases around an equilibrium position; torsion and flexural oscillations of the double helix (10−10–10−8 s); large-amplitude movements of phosphates, sugars and bases occurring at the transition of DNA double helix from

Interactions of Metal Ions with DNA 33

one form to another (10−7–10−5 s).34 Thus for modelling of DNA structural changes stimulated by interactions with various transition metal ions it is necessary to estimate the correlation between the energy of interaction and the lifetime of the complexes formed.

2.2 Interactions of Metal Ions with DNA 2.2.1 Thermodynamic Adsorption Model of Complex Formation The catalytic characteristics of DNA are closely related to structural changes caused by interactions with different species, particularly with metal ions. Evidently, transition metal ions can locally influence the structure of DNA only when the lifetimes of the complexes are commensurable with the specific times of inner mobility in DNA (oscillation of small groups of atoms, untwisting of the double helix, opening of the individual base pairs, untwisting of helix binding of proteins and cell division). The movements last from 10−10 s to hundreds of seconds and longer. We have managed to find a correlation between dynamics and stability28 by applying Frenkel’s phenomenological thermodynamic approach,35–37 which was used as early as 1924 for the study of gas adsorption on the surface of solids. Frenkel introduced a new term ‘lifetime’ (t) for the adsorption state and connected it to the energy of interaction |∆E| between adsorbate and adsorbent surface by the following expression:

τ = τ 0a−s ⋅ exp ( ∆E kBT )

(2.1)

where τ 0a−s is the time of a single oscillation of the adsorbate on the surface, assumed to be 10−13–10−12 s and kB is the Boltzmann constant.36 The above model can, with good approximation, be applied to the study of interactions between small molecular species and biomacromolecules in solution. Using the relationship DG = −RT lnK, a simplified expression of (2.1) can be derived:

τ = τ0 ⋅ K

(2.2)

where t is the lifetime of a metal ion–macromolecule complex. Now we must clarify the value t0 for solutions. Frenkel’s τ 0a−s is the duration of the fluctuation excitation of adsorbing atoms or molecules interacting with a solid surface. It is supposed to be equal to the period of oscillation of the adsorbate relative to the adsorbent surface. In solutions, t0 describes the duration of relaxation of rotary and translation movements of the solvent molecules, ions, solvated ions or low-molecular-weight substances and lies between 10−11and 10−10 s. Thus, if we assume a pK in the range 4–6 for binding of DNA with two-fold positively charged metal ions of the first transition series, and t0 = 10−11 s, then the life span of these complexes is in the range 10−7–10−5 s.

34

Thermodynamic Models

The expression t = t0 · K allows us to correlate the stability constant of complex formation to the lifetime of the complex. Thus, the principal concept of molecular biophysics regarding biomolecules: structure – dynamics – function can be reformulated as: structure – stability – function. One may notice that such an approach highly simplifies and widens the time interval (from 10−10 s to 105–106 s and more) for investigating dynamic characteristics of macromolecules. A detailed description of the derivation of formulae (2.2) is given in reference28. 2.2.2 Stability Constants of DNA–M2+ Complexes Stability Constants Determined by Graphical Methods Based on Adsorption Isotherms Different methods suitable for calculation of stability constants have been developed, for the most part based on linearization procedures of the law of mass action.38,39 The improvement of computer technology led to the development of nonlinear curve fitting algorithms;40,41 among these the Scatchard and Klotz methods have been widely used to analyze experimental data.42,43 Let us describe, as an example, various models of Pb2+ binding to DNA using modified Scatchard plots and two types of the Klotz graphical representation. Figure 2.1A shows the Scatchard plot. The intersection of the curves with the abscissa gives the value n = n1 + n2 (n is the number of binding sites for Pb2+ per phosphate group of DNA at saturation). The intersection of the curves with the ordinate gives the value of the binding constant (K): Kn = k1n1 + k2n2.43 The parameters are calculated by the nonlinear least-squares fit to the equations: r m = 0.5  B ( r ) + B2 ( r ) + 4C ( r )  B ( r ) = k1n1 + k2 n2 − ( k1 + k2 ) r C ( r ) = k1k2 r ( n1 + n2 − r ) Figure 2.1B illustrates the Klotz method with intercepts a1, a2, a3, a4 for variables r/m and r, where two independent classes of binding sites are assumed. a1 = n1k1 + n2 k2 a = n + n 1 2  2 System:  2 2 2 a3 ( n1k1 + n2 k2 ) = a1 ( n1k2 + n2 k1 ) a4 = a2 2 k1k2 It is seen from Table 2.244 that the parameters determined by the Klotz and modified Scatchard analysis methods are almost identical. Consequently, it can be assumed that both models can be used to determine binding constants of Pb2+ to DNA. Based on the c2 test, the Scatchard method (c2 = 0.81) seems to give a better estimate than the Klotz method (c2 = 3.08).45

Interactions of Metal Ions with DNA 35 1.2

1.2

1.0

1.0

Pb(II)

A

0.6

0.6 0.4

0.4

0.2

0.2 0.0 0.0

B

0.8 r/m × 104

r/m × 104

0.8

a1

a4 a3

0.0 0.1

0.2

r

0.3

0.4

0.5

0.0

0.1

a2 0.2

r

0.3

0.4

0.5

Figure 2.1A: The Scatchard plot for binding of Pb(II) with DNA (2 mM NaNO3, 20 °C) as non-linear least square fit r m = 0.5 B (r ) + B2(r ) + 4C (r )  , where B(r) = k1n1 + k2n2 − (k1 + k2)r and C(r) = k1k2r(n1 + n2 − r). Figure 2.1B: The Klotz plot for binding of Pb(II) with DNA (2 mM NaNO3, 20 °C). Calculated values (k1, n1, k2, n2) were obtained from the Klotz equation system.

Table 2.2 Parameters of Pb2+ binding with DNA in 2 mM NaNO3, 20 °C Pb

Stability constant K × 104, M−1 Free Gibbs energy −∆G0, kcal mol−1 Number of binding sites at saturation n Microconstant k1 × 104, M−1 Number of binding sites n1 Microconstant k2 × 104, M−1 Number of binding sites n2 c2 distribution

Analysis by the Scatchard method

Analysis by the Klotz method

2.4 ± 0.5 5.96 0.48 ± 0.05 21.1 ± 0.7 0.043 ± 0.002 0.56 ± 0.01 0.44 ± 0.03 0.81

2.3 ± 0.2 5.94 0.50 ± 0.06 24.7 ± 1.2 0.034 ± 0.001 0.68 ± 0.01 0.47 ± 0.01 3.08

The stability constants (log K) for Mg2+, Cr3+, Mn2+, Co2+, Ni2+, Cu2+, Zn2+, Ag+, Cd and Pb2+ binding to DNA are in the range 4–6 (Table 2.3), and assuming a t0 value of 10−11 s, the lifetimes t fall in the range 10−7–10−5 s. For DNA this time corresponds to the movements of phosphates, sugars and bases with large amplitudes (see introduction). These movements are related to conformational changes and untwisting of the double helix. As metal ions of the first transition series form chelates with DNA, the stabilities of the complexes may be estimated from the DNA 2+

36

Thermodynamic Models

Table 2.3 Binding constants of metal ions with ctDNA (20 mM Na+). Metal ions

Cr3+

Co2+

Ni2+

Cu2+

Zn2+

Ag+

Cd2+

Pb2+

Log K

3.85

4.3

4.64

5.1

4.2

5.84

4.1

4.25

Ag+ Pb2+

1.0

Cu2+ Ni2+ Co2+ −Cr3+ Zn2+ Cd2+ H+

0.9

LogK

0.8 0.7 0.6

Mg2+ 0.5 ϕ

0.4 M-DNA 0.3 −3.0

−2.5

−2.0

−1.5

−1.0

−Log[Na(1)]

Figure 2.2 Efficacy of Na+ concentration on the stability constants (log K) for ctDNA complexes with Mg2+ 43 Co2+, Ni2+, Cu2+, Zn2+, Cd2+, Pb2+, Cr3+, Ag+ and ϕ, which is the electrostatic potential of a cylindrical polyion surface (r = 1 nm) obtained from the Poisson–Boltzman equation44 reduced to unity

dynamics. For example, the difference in pK between Co2+ and Cu2+ binding to ctDNA is only 0.8 (Table 2.3) compared to the much larger differences for binding of Co2+ and Cu2+ to bidentate ligands like NH2CH2CO2− and ethylendiamine (en) where the pK varies from 4.6 to 8.6 and from 6 to 11, respectively.46 Thus, when M2+ ions interact with DNA, the dynamic properties of DNA connected with big amplitude movements, e.g. unwinding the double helix and opening of base pairs, determine to a large extent the stability constants for complex formation. Effect of Ionic Strength Figure 2.2 is a graphic representation showing how pK varies with ionic strength for H+ and metal ions (Cr3+, Co2+, Ni2+, Cu2+, Zn2+, Ag+, Cd2+, Pb2+) binding to DNA. An NaCl salt solution was used for all metal ions except Ag+ and Pb2+ where NaNO3 was used. For Mg2+ the binding constants with poly(A-U) are given in reference.47

Interactions of Metal Ions with DNA 37

In addition, the dependence of the surface electrostatic potential (ϕ) of a cylindrical polyion (r = 1 nm) is plotted as a function of the ionic strength of the solution as obtained from the Poisson–Boltzmann equation.48 For Mg2+ the pK dependence and the surface potential are closely related, indicating a nearly exclusive binding of Mg2+ to the negatively charged phosphate groups of DNA. One may notice that the binding constants of Ag+ and Pb2+ are only weakly dependent on ionic strength. This is in agreement with the expected inner-sphere complexation of Ag+ ions involving nitrogen atoms of the nucleobases (soft binding). Divalent lead is one of the post-transition metal elements that exhibits the so-called ‘inert-pair effect’. This lone pair can cause a non-spherical charge distribution around the Pb2+ cation. ‘Stereochemically active lone pairs’ can take various forms in Pb2+ compounds.49 Recent data show that Pb2+ forms hemidirect coordination compounds both in complexes with water and in complexes with O and N atoms of ligands,50 which, in turn, are reactive centres in DNA for other metal ions. The stereochemistry of Pb2+ may be the reason of such a low-effect action of ionic strength. The log K dependencies for the transition metal ions Cr2+, Co2+, Ni2+, Cu2+ and 2+ Zn are in between the two extremes mentioned above. The slopes of their lines allow determination of the ‘softness’, or ‘transition degree’, of these ions assuming that the slope angle of the lines is proportional to this property. Effect of DNA G-C Content on Metal Complexation To quantify the effect of DNA heterogeneity upon metal ion binding, the values of pK for Ni2+ complexes with DNA of various origins (T7 phage, calf thymus, bovine spleen, salmon sperm, Spirulina platensis, E.coli) have been determined.51,52 In Figure 2.3

5.4 Ni(II)-DNA

E.coli T7

5.2 Spirulina platensis LogK

5.0

4.8 Bovine spleen

Calf thymus Salmon sperm Mice liver

4.6

40

42

44

46

48

50

52

(GC) %

Figure 2.3 Dependence of Log K versus G-C content

38

Thermodynamic Models 4.0 Ag+

M-DNA 20 mM Na+

3.5 Pb2+ 3.0 X m2 r

Cu2+ Co2+

2.5

2.0

Cd2+

Ni2+ y = −1.31 − 0.85x; R 0.92 SD 0.05; N7; p < 0.0001;

1.5 −6.0

−5.5

−5.0 pK

Zn2+ Cr3+ −4.5

−4.0

−3.5

Figure 2.4 Correlation between the covalence index Xm2r and − log K = pK for M–DNA complexes

pK for Ni2+ binding to different species is shown to increase with G-C content. The empirical dependence of pK versus G-C content has been derived: pK = 1.96 + 0.07 (% G-C). The observed trend to form stronger complexes with increasing G-C content may be due to the high probability of G-N7/Ni2+ complex formation. A linear correlation has been observed between log metal/poly d(GC), assessed at the midpoint of the structural transition curve, and the covalent index ( X m2 r ) 53,54 (Xm being the Pauling electronegativity and r the effective ionic radius55). The authors propose that the mid-phase mole ratio is inversely proportional to the stability constant for the metal ion–polydeoxynucleotide interaction. Such dependence between binding constants and X m2 r was also obtained in our case. The linear correlation of pK (for complexes of metal–ctDNA) and the covalent index X m2 r presented in Figure 2.4 suggests that the covalent contributions to the overall interaction energy are significant and demonstrates the dynamic nature of metal–DNA interactions (except in the case of Pb2+). These results are in good agreement with data in references 53 and 54. 2.2.3 UV Spectroscopic Study of Proton and Metal–DNA Interactions Table 2.4 shows spectral changes of UV absorption spectra of ctDNA and that of poly(dG-dC) · poly(dG-dC) caused by different ligands and macroscopic changes of the surrounding medium. Dn in the table is calculated from the expression Dn = Des/K, where Des = |De(nmax)| + |De(nmin)| is the total intensity of ultraviolet difference spectrum, K = |e(1)(nmax)| + |e(1)(nmin)|; e(1) is the first derivative of e(ν) function. Dn for d(pG) + d(pC) + H3O+ (pH = 2), H3O+, Cu+ and Ag+ were determined by absorption spectra of DNA complexes (more precisely this issue is observed in reference 17). Analyzing this acid bathochromic shift (the first line in the table) and

Interactions of Metal Ions with DNA 39 Table 2.4 Bathochromic shift (∆u) of UV absorption spectra of ctDNA and poly(dG-dC)·poly(dGdC) caused by different metal ions and H3O+ Ions

d(pG) + d(pC) + H3O+ H3O+ Mg2+ Mn2+ Co2+ Ni2+ Ni2+ + 40% ethanol Ni[(en)(H2O)24+ ] Ni[(en)(NO2−)(NCS−)]0 Ni2+ + EtBr Z-DNA poly(dGdC)·poly(dG-dC) 25% ethanol, 102 NaCl, (0.75 Ni2+/P) Cu2+ Cu+ Zn2+ Ag+

Bathochromic shift of the absorption band (∆u(cm−1)) ∼700 295 15 65 110 150 185 75 35 180 ∼700

190 ∼700 85 ∼500

Change of the Widening of the DPT formation absorption absorption band energy in G-C pair U intensity (%) (cm−1) and special characteristics (kcal mol−1) Insignificant 2.5↓

Insignificant

– 0.5 2.4 1.4 1.1 0.9

140 (1/2h)

0.8 0 1.3 0.2

1.5↓

7.4↓ 10↓

1000 (2/3h)

taking into account absorption of adenine and thymine, the amount of which totals 60% in ctDNA, we can roughly accept that the joint effect of the shift from protonation of guanine and cytosine is ∼700 cm−1 per 1 mol of G-C pair. Correlation Between Du and Ke/aq Interactions of aqua ions with the DNA double helix have a discrete nature due to steric requirements characterized as inner-sphere, outer-sphere or double outersphere interactions. The latter refers to interaction of hexa-aqua ions with the hydration layer of DNA. One may also take into account so-called ‘atmospheric’ interactions which involve mobile adsorption within the effective diameter of DNA depending on the ionic strength.56 UV spectroscopy is a sensitive method for detection and registration of the mentioned interactions. As for thermodynamic methods, they reflect the integral characteristics of these interactions. That is why stability constants (pK) are less dependent on G-C composition, ionic strength and the type of ions. The soft structures of transition metal–DNA interactions are also caused by the fact that each of the six water molecules surrounding the ions may exist in two positions (two nonbonding pairs of electrons in the water molecule are localized on ′ ).57 the sp3 hybrid orbitals of the oxygen and the distance between them is ~ 0.6 Å Thus, the water molecules located in a six-coordinated sphere of hydration can exist in 26 different states.

40

Thermodynamic Models 500

Ag+

400

*Cu+ H+

∆ν

300 Cu2+

Ni2+ + ethanol

200

100

Mn2+ Mg2+

0 0.0

Ni2+

Zn2+ Co2+

Ni(en)2+

Ni[(en)(NO2)(NCS−)]0 0.4

0.8

1.2

1.6

2.0

1/∆Ga

Figure 2.5 Correlation between ∆ν of ctDNA with different metal ions and 1/∆Ga. ∆Ga refers to the reaction of the hydrated electron with the ion under study and is related to the rate ∆Ga . The vertical factor for Cu+ is ×2 constant by the expression K e aq = 1011 exp − RT

(

)

More details follow from Figure 2.5, which exhibits the correlation between the shift of the absorption band (Dn) of DNA bound to the investigated ions and the reaction rate constants (Ke/aq) of hydrated electrons with the same ions, particularly with the 1/DGa values (Table 2.1). DGa is the hydrated electron activation energy in the expression: Ke/aq = 1011exp(−DGa/RT).29,30 At I = 0.01 M, a linear relationship between these values seems to exist only for Mn2+, Co2+, Ni2+ and Zn2+ while Mg2+, 0 Cu2+, H+, Ag+ and Cu+ deviate from the line. So do Ni(en)2+, Ni [( en)2( NO−2 )( NCS− )] 2+ and Ni (40% ethanol). From the UV, visible and near IR spectroscopic studies it may be concluded that Mn2+, Co2+, Ni+2 and Zn+2 form outer-sphere complexes with DNA. The small influence of these divalent ions on the DNA absorption spectrum and the correlation of Dn with 1/DG suggest that their interactions with the DNA nucleobases in solution take place through hydrogen bonding involving their hydrated coating. A 31P NMR spectroscopy study of the interaction between DNA and the paramagnetic ions Mn2+ and Co2+ showed that only 15 ± 5% of the total bound ions was directly coordinated to the phosphate groups (inner-sphere complexes). The remaining Mn2+ and Co2+ ions were bound either as outer-sphere complexes relative to the phosphate groups or elsewhere to the DNA, possibly to the bases.58 These results are in agreement with X-ray crystallographic data, which show that transition metals may coordinate to mononucleotides by direct nucleobase Nbinding and/or phosphate binding.59,60 The UV data indicate that Ag+, H+, Cu+ and, partially, Cu2+, interact directly with DNA without participation of water molecules. 0 Ions of Mg2+, Ni(en)2+ and Ni [( en)2( NO−2 )( NCS− )] , which stay below the line (Figure 2.5), have the lowest affinity to DNA nucleobases. In the case of direct binding of metal to nucleobases in DNA there is a certain probability of electron tunnelling transitions from the nitrogen bases to the metal

Interactions of Metal Ions with DNA 41

ion. As an example, for the DNA–Cu2+ complex a total electron transition of 0.2 quantum output is observed under UV irradiation.31 Thus, H+, Cu2+, Cu+ and Ag+ can form inner-sphere interactions with DNA while the barrier mechanism is typical for Mn2+, Co2+, Ni2+ and Zn2+. Hence, we may conclude that ions such as Mn2+, Co2+, Ni2+ and Zn2+, as well as most of the Cu2+ ions form outer-sphere complexes with DNA. Experiments with Ni2+ indicate that these ions prefer the GC/CG and CG/GC type base pairs causing a red shift of the DNA absorption spectrum. Influence of GC Composition The influence of the DNA nucleotide composition on the ultraviolet difference spectrum (UDS) of DNA–metal ion complexes has been investigated for Co2+, Ni2+, Cu2+ and Zn2+. Most attention has been paid to Ni2+ and its interactions with DNA of various origins: Clostridium perfringens (27% GC), mouse liver (C3HA line, 40% GC), calf thymus (40% GC), salmon sperm (44% GC), herring sperm (44% GC), E. coli (51% GC), Micrococcus luteus (72% GC). In addition, interaction of Ni+2 with the oligonucleotides poly(dG-dC).poly(dG-dC), poly(dG).poly(dC) and poly(dA-dT).poly(dA-dT) have been studied. These results are shown in Figure 2.6 where Des versus the GC percentage is plotted. The Des values are given for the relative concentration of 0.25 Ni+2 per nucleotide unit. Analysis of the data for the interaction of Ni+2 with DNA shows preferential binding to GC pairs. Ni2+-Induced B → Z Transition in poly(dG-dC) · poly(dG-dC) Polynucleotide Figure 2.7 presents the absorption spectra of poly(dG-dC)·poly(dG-dC) in the presence of 0.25% ethanol solution (10−2 M NaCl without (1) and with (2) Ni2+ ions at

600 Ni2+

Poly(dG-dC)Poly(dG-dC)

500 400 ∆εs

E.coli Micrococus luteus

Herring sperm Salmon sperm Calf thymus

300 200

Mice liver(C3HA line) Poly(dG)poly(dC) Cl.perfringers

100 0 −20

Poly(dA-dT)Poly(dA-dT) 0

20

40

60

80

100

GC% +2

Figure 2.6 Influence of Ni on the UV spectra of various DNAs and polynucleotides. Graphical illustration of the dependence of the mole extinction (∆es) on the percentage of GC pairs present

42

Thermodynamic Models

0.8

1 2

A

0.0 45 000

36 000 ∼ ν, cm−1

Figure 2.7 UV spectra of polynucleotide poly(G-C).poly(G-C) in 25% ethanol solution (10−2 M NaCl) without (1) and with (2) Ni2+ ions; concentration is 0.75 Ni/P

5 4 3 1000

2

∆ε

0 1 − 1000

6 7

−2000 45 000

40 000

35 000 ∼ ν, cm−1

Figure 2.8 UDS of polynucleotide poly(G-C).poly(G-C) in 25% ethanol solution at different concentrations of Ni2+: (1) 0.1 Ni/P, (2) 0.25 Ni/P, (3) 0.3 Ni/P, (4) 0.4 Ni/P, (5) 0.5 Ni/P, (6) 0.75 Ni/P, (7) 1 Ni/P

concentrations of 0.75 Ni2+ per nucleotide (P)). Ni2+ ions cause a significant red shift of 700 cm−1 and strong hypochromic effect (25%). Figure 2.8 shows the UDS of poly(dG-dC)·poly(dG-dC) in 0.25% ethanol solution with Ni2+/P ratios in the range 0.1 to 1.0. Figure 2.9 demonstrates a large difference between the UDS of Ni2+/ poly(dG-dC)·poly(dG-dC) recorded in water (1) and in 25% ethanol solution (2). In the first case a typical saturation line is shown while in the second case an S shape curve characteristic of conformational change is observed, e.g. B→A or B→Z transition. The CD spectra (Figure 2.10) recorded under the same conditions as for UDS show that at concentration Ni/P = 0.2 the polynucleotide is in the C conformation (commonly observed as a low humidity form of the lithium salt). At higher concentrations of Ni+2, the double helix is shown to undergo transition to the

Model and Mechanisms for Point Defects 43 3250

∆εs

2

1

0

1 Ni(II)/P

Figure 2.9 The dependence of UDS of polynucleotide poly(G-C).poly(G-C) on Ni2+ concentration in water (1) and in 25% ethanol solution (2)

7 6 4

1

εL − εR

2

3

5 4 −8 40 000

∼ ν, cm−1

32 000

Figure 2.10 CD spectra of polynucleotide poly(G-C).poly(G-C) in 25% ethanol solution at different concentrations of Ni2+: (1) without Ni2+, (2) 0.1 Ni/P, (3) 0.2 Ni/P, (4) 0.25 Ni/P, (5) 0.3 Ni/P, (6) 0.35 Ni/P, (7) 0.4 Ni/P

Z conformation. It was also demonstrated in our experiments that the Ni2+–poly(dGdC)·poly(dG-dC) complex forms the C conformation with wrong base pairs during the B→Z transition process.61–63 Thus, we have demonstrated that UV spectroscopy is a sensitive method for monitoring conformational transitions in double-helical DNA.

2.3 Model and Mechanisms of Metal-Induced Formation of Point Defects 2.3.1 Double Proton Transfer (DPT) in DNA: UV Spectroscopic Measurements The influence of pH on UV spectra of guanine and cytosine, as well as on keto–enol and amino-imino transformations in guanine and cytosine, has been known for a

44

Thermodynamic Models

long time.46,63,64 It should be noted that the transfers of protons between the donor and acceptor groups N1 and O6 in guanine and N4 and N3 in cytosine are definitely performed with the help of the hydrated water molecules; they form a kind of cyclic structure with the above groups in guanine and cytosine. Because of self-congruent transition of protons from N1 of guanine and N4 of cytosine to molecules of hydrated water, reorientation of water molecules takes place in both cases. Thus, we can state that the specific energy of H-bonds with two energy states and self-congruent transfer of protons in cyclic structures is a necessary and sufficient condition for keto–enol and amino–imine tautomeric transformations of polar organic molecules, particularly guanine, cytosine and adenine in solution. Water molecules act as a mediator in the process of forming the joint electron–proton complementary complex between the proton-acceptor/donor groups of the solute molecules and those of the solvent. As already mentioned, there is no need for water molecules as mediators in the DNA duplex because a pair of complementary bases always forms a cyclic structure. Double proton transfer (DPT) in DNA is demonstrated spectroscopically as bathochromic shifts, weak hypochromic effects and small widening of the absorption band (Table 2.4).14,17 We shall discuss the mechanism in detail, using GC pairs as an example for the following reasons: (i)

(ii) (iii) (iv) (v)

the effect of H+ interactions with guanine and cytosine,47,64 as well as interaction between transition metal ions and guanine are easily observed in UV-spectra; 65 spontaneous mutations of a genome occur more often at GC pairs than at AT pairs;66–68 guanine and cytosine are often populated with rare enol and imino forms;69–71 GC pairs are far less resistant to tunnelling transitions compared to AT pairs; 26,72,73 in the DNA duplex the site of preferable binding of H+ and metal ions is the endocyclic N7 of guanine located in the major groove.28,74

In general, the keto–enol and amino–imine tautomeric transformations in GC pairs are conditioned by the electric charges on G-N1 and C-N3 of endocyclic nitrogens, which play the principal role in H-bonding of the base pairs. Figures 2.11 and 2.12 present electronic configurations of atoms participating in the formation of Hbonds in GC and AT pairs. It is interesting to consider here the disturbance of the electronic structure of guanine in the pair of bases in a DNA duplex provoked by H+ and transition metal ions. When a positive ion interacts with a pyridine-type nitrogen (G-N7), a nonbonded pair of electrons located in a sp2 hybrid orbital can significantly overlap the p-electronic system of the indole ring of guanine inducing a decrease of electronic density of the ring, including the endocyclic N1 nitrogen atom. This will lead to a decrease in the depth of potential energy from the N1 side of guanine and an increase of possible proton tunnelling to N3 in cytosine, as illustrated in Figure 2.13. As early as in 1969, Scheiber and Daune47 pointed out quantitative similarities

Model and Mechanisms for Point Defects 45

Guanine

Watson-Crick base pair Cytosine

H

N2 pz

e

N1 sp

pz

sp2

O6 pz

sp2 N2

pz

O2

e

e H

H e

e

H wrong pair

sp2

N1 pz

2

sp2

O6

Watson-Crick base pair

sp N 3

Adenine py pz

sp2N

e

pz

sp2

sp2 N 1 1

e

4

sp2

H

sp2

sp2

O4

H e

e

2pz

H

N6

sp2 N

pz

sp N

pz 3

e

sp2

pz

spz

H

1

O4

N3 sp2

N4 pz

pz

H

wrong pair

O2

p pz sp N y 3

H

pz

e

2pz

sp2

pz

Thymine e

N6

sp2

sp2

pz

A

pz

B

Figure 2.11 The electron configuration of atoms of G-C (A) and A-T (B) pairs taking part in H-bonds before and after DPT

minor groove N2

G

N1

e N7 Mn+

O6

H

O2

H

N3 H

N4

e C

major groove

Figure 2.12 Double proton transfer (DPT) in a G-C pair of DNA

E 0

r

+

N

H

N

Figure 2.13 Hypothetical function of potential energy of N-H···N type H-bonds

46

Thermodynamic Models

between difference spectra of G-C + H+ and DNA + M2+ (M2+ = Cu2+, Ni2+ and Fe2+). Our investigations, carried out on metal ion–DNA complexes by thermodynamic and spectroscopic methods, in particular spectrophotometry in the UV, visible and near IR ranges, allowed us to propose a scheme for the structure and energetics of DNA interactions with metal ions.31,75 2.3.2 Depurination Interaction of H+ and Mn+ with N3 and N7 of guanine in DNA may induce DPT (1) and/or hydrolysis of the glycosidic linkage C1–N9 (2): C1+ C1+ + OH−

N9−

and

C1 OH

and

N9− + H+

(1)

N9H

(2)

The process of depurination may take place only in the unwound state of double-helical DNA. Crothers and collaborators76 studied the energetics of the opening of the central base pairs in different triplets of double-stranded RNA, which was often used as a model for the DNA double helix since crystallographic data for DNA was not known at the time. According to this study the energy of opening of a G-C pair at 25 °C is at a maximum and equals 7.5 kcal mol−1 in the sequence: G G G C C C corresponding to a minimum probability of 0.3 × 10−5 s, while the energy of opening an A-U pair is at a minimum (4 kcal mol−1) for the sequence: A A A U U U corresponding to a maximum probability of 120 × 10−5 s. By inserting these data in Equations (2.1) and (2.2), we obtain the times ∼3 × 10−6 s for G-C and ∼10−8 s for A-U. The probability of metal-induced DPT in DNA double helix is equal to the ratio of the wave shift caused by metal ion interactions with DNA G-C pairs to the wave shift after total protonation of guanine (pH 1) and cytosine (pH 2), which amounts to ∼700 cm−1:15 PDPT = ∆vMn+ ∆vG+C+H+ = ∆vMn + 700 cm −1 Considering the values of Dn from Table 2.4, we can assume that for the metal–DNA complexes studied by us these probabilities are in the range from 10−1

Model and Mechanisms for Point Defects 47

(for Mn2+) to 1 (for Cu+).15 Consequently, metal-induced depurination of the DNA chain in wrong Watson–Crick pairs depends mainly on the probability of base pair opening. In conclusion, interactions of H+ and Mn+ with N3 and N7 of purines will increase the probability of prototropic tautomerization of purines and, finally lead to the transition of N9 purines in pyrol-type nitrogens followed by consecutive depurination. Later the question of base pair opening was studied directly in DNA under different conditions using NMR and other methods.77–81 The results obtained by these authors are in agreement with the model proposed by Crothers.76 2.3.3 Formation of Point Defects For steric reasons, metal-ion-induced DPT may take place only at G-N7 in GC pairs, which can be approached by transition metal ions in DNA. DPT may also take place at G-N3 (GC pair) and T-O2 (AT pair) sites, but in this case only H+ ions can approach these sites. Thus, it may be concluded that transition metal ions cause DPT exclusively in GC pairs via G-N7, while H+ can do it at both GC and AT via G-N7, G-N3 and T-O2. DPT and depurination in vivo can lead to the gene mutations i.e. point mutations (transition and deletion, respectively). In the first case, the geometry of the base pairs that undergo DPT remains the same, or changes very slightly, and it may stay unnoticed in the processes of replication and correction. Nevertheless, the hydration of the correct Watson–Crick pairs differs from the wrong ones (transferred by DPT).14 We do not know the biological mechanisms of detection of these differences. In the view of solid-state physics, one of the major characteristics of these kinds of defect is their formation energy (U). In the case of DNA, this energy lies in the range 0 (for Cu+) to 2.4 kcal mol−1 (for Mg2+) (Table 2.4). The values of the energy (U) given in the table were obtained by the Boltzman distribution. In our case, PDPT = ∆vMn + 700 cm −1 represents the ratio of G-C pairs in keto and enol tautomeric forms: PDPT = nket/nen ≈ e−U/RT. Let us consider the probable mechanism of transition-type mutations in a G-C pair of DNA by an example involving the interaction of Cu2+ with G-N7. Schematically we can describe it as follows: the helix unwinding

Cu2+

G DPT G* C

C*

G*

mutation A

C

T

A*

A

C

T

* – rare tautomeric form of bases

48

Thermodynamic Models

Cytosine can stay in its imine form for quite a long period of time, in spite of the fact that this structure is energetically less advantageous compared to that of the amine form. The restoration process includes the time needed for the formation of a cyclic unidirectional structure of water molecules. Thus, when a wrong G-C pair opening takes place, the less hydrated cytosine needs to pass through the potential barrier in order to rebuild the surrounding water in a unidirectional cyclic structure. The time span depends on the number of water molecules participating in the cycle and in many cases exceeds the time needed for the closing of the helix. In the process of winding/unwinding of the double helix in the presence of transition metal ions, one may envisage the following alterations: (1) Mn+

G* C*

(2) M

n+

G* C

(3) Mn+

C*

G C*

(4) Mn+

G C

For the second and third examples, the defect that remains after closing of the disrupted pair is easily recognized by the repair system in vivo. In particular, one should notice that Cu2+ ions can easily be reduced to the Cu+ state (UV irradiation, g irradiation, ascorbic acid etc.)28,31 and the Cu+–DNA lifetime will increase by about 11 orders of magnitude, reaching 104 − 5 × 105 s.17 In this case, Cu2+, which is essential for life for many mammals, can be transferred into extremely toxic soft Cu+ ions. Depurination is the second process inducing point defects. Interaction of H+ with pyridine-type nitrogen atoms, and the preference of transition metal ions to bind to G-N7 may lead to the hydrolysis of the C-N3 glycosidic linkage, which is the weakest covalent bond in DNA with a torsion rotation barrier in the range 0.003–0.006 kcal mol−1 74,82 At the moment of base pair opening, the ions will initiate rupture of the bond and if it is not restored in vivo, severe damage of the double helix called deletion will take place. Even if the repair system restores the deleted base, this may not completely prevent damage, because interaction of metal ions with the DNA duplex may also lead to DPT (see above), and after depurination of guanine, cytosine adopts a rare tautomeric imine form. In this case, adenine instead of guanine will be inserted in the base pair.

2.4 Conclusions (i)

A thermodynamic model of metal ion interactions with DNA can be used to establish direct proportional correlation between dynamics and stability constants. This approach is based on the physical model of gas and vapour adsorption on the surface of solids. Thus, the principal concept of molecular biophysics regarding biomolecular structure–dynamics–function can be reformulated as structure–stability–function. One may notice that such an approach highly simplifies and widens the time intervals (from 10−10 s to 105–106 s and more) under investigation for dynamic characteristics of macromolecules.

Abbreviations

49

(ii) The dynamic properties of DNA connected with big amplitude movements like unwinding of the double helix and opening of base pairs determine the values of the stability constants of DNA complexes with transition metal ions, as well as the ranges of their variations. (iii) UV spectroscopic manifestation of double proton transfer (DPT) in GC pairs induced by H+ ions and transition metal ions, and the mechanisms of this phenomenon can be considered using a phenomenological quantum-mechanical approach to assess the effect of environment change on electron configuration of atoms in molecules participating in H-bonds. This consideration takes into account the specific energy of double-level H-bonds, and self-congruent transition of protons in cyclic structures. These structures form unified electronic– proton complementary molecular complexes between proton-donor and proton-acceptor groups. The latter is an indispensable and satisfactory condition for keto–enol and amino–imino tautomeric transformations in solutions of polar organic molecules. (iv) A comparison of the Scatchard plot and the Klorz method for determining stability constants, using, for example, the Pb2+/DNA interaction, shows that the former method gives the best results. (v) The use of UV spectroscopy has demonstrated that Ni2+ ions under relatively mild conditions (25% ethanol, 0.75 Ni2+/DNA(P), 10−2 M NaCl) induce B ↔ Z transitions in double-helical DNA. (vi) A mechanism for point defect formation induced by metal ions, leading to mutations of transition or deletion types in DNA, has been presented, using as an example the interaction of Cu2+ ions with DNA. It was shown that formation of metal-induced wrong Watson–Crick base pairs via DPT in water solutions requires energy within the range 0 (for Cu+) to 2.4 kcal mol−1 (for Mg2+).

Acknowledgements The authors express their gratitude to Prof. J. Monaselidze for useful discussions; Dr M. Kharatishvili for providing the CD spectra of DNA; Dr I. Khutsishvili, Dr E. Mchedlishvili and Dr Kh. Sologashvili for their active assistance in this work. The work was partly supported by the grant number 2.29.04 of the Georgian Academy of Sciences.

Abbreviations ctDNA Calf thymus DNA CD Circular dichroism DPT Double proton transfer e Molar extinction coefficient Des = |De(nmax)| + |De(nmin)|

50

Thermodynamic Models

EtBr En − eaq Ke/aq K IR M UDS

Ethidium bromide Ethylenediamine Hydrated electron − Rate constant of the reaction of eaq Stability constant Infrared Free hydrated metal ion Ultraviolet difference spectra

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17. Bregadze, V.G.; Khutsishvili, I.G.; Chkhaberidze, J.G.; Sologashvili, K.; DNA as a mediator for proton, electron and energy transfer induced by metal ions; Inorg. Chim. Acta, 2002, 339, 145–159. 18. Bregadze, V.G.; Chkhaberidze, J.G.; Khutsishvili, I.G.; Effect of metal ions on the fluorescence of dyes bound to DNA. In: Metal Ions in Biological Systems, Vol. 33, A. Sigel and H. Sigel, Eds., Marcel Dekker Inc., New York, Basel, 1996, Chap. 8, 253–266. 19. Chalote Auerbach, F.R.S.; Mutation Research: Problems, Results and Perspectives. Mir, Moscow, 1978. 20. Verne, G.; The Evolutionary Process: A Critical Review of Evolutionary Theory, Mir, Moscow, 1985. 21. Bloemink, M.J.; Reedijk, J.; Cisplatin and derived anticancer drugs: mechanism and current status of DNA binding. In: Metal Ions in Biological Systems, Vol. 32, A. Sigel and H. Sigel, Eds., Marcel Dekker Inc., New York, Basel, 1996, Chap. 19, 641–675. 22. Vinje, J.; Sletten, E.; NMR spectroscopy of anticancer platinum drugs; Anti-Cancer Agents Med. Chem., 2007, 7, 35–54. 23. Liu, Y.; Sivo, M.F.; Natile, G.; Sletten, E.; Antitumor trans-platinum complexes can form cross-links with adjacent purines; Angw. Chem. Int. Ed., 2001, 40, 1226–1228. 24. Liu, Y.; Vinje, J.; Pacifico, C.; Natile, G.; Sletten, E.; Formation of adenine-N3/guanine-N7 cross-link in the teaction of trans-oriented platinum substrates with dinucleotides; J. Am. Chem. Soc., 2002, 124, 12854–12862. 25. Karidi, K.; Garoufis, A.; Tsipis, A; Hadjiliadis, N.; Dendulk, H.; Reedijk, J.; Synthesis, characterization, in vitro antitumor activity, DNA binding properties and electronic structure (DFT) of the new complex cis-(Cl,Cl)[RuCl2(terpy)(NO)]Cl; J. Chem. Soc. Dalton, 2005, 1176. 26. Löwdin, P.O.; Proton tunneling in DNA and its biological implications; Rev. Mol. Phys., 1963, 35, 724–732. 27. Löwdin, P.O.; Quantum genetics and the aperiodic solid. Some aspects of the biological problems of heredity, mutations, aging and tumors in view of the quantum theory of the DNA molecule; Adv. Quant. Chem., 1966, 2, 213. 28. Bregadze, V.G.; Metal ion interactions with DNA: consideration on structure, stability, and effects from metal ion binding. In: Metal Ions in Biological Systems, Vol. 32, A. Sigel and H. Sigel, Eds., Marcel Dekker, New York, 1996, Chap. 12, 419–451. 29. Anbar M.; Neta P.; A compilation of specific bimolecular rate constants for the reactions of hydrated electrons, hydrogen atoms and hydroxyl radicals with inorganic and organic compounds in aqueous solutions; Int. J. Appl. Radiat. Isotopes, 1967, 18, 493–523. 30. Hart, E.J.; Anbar, M.; The Hydrated Electron, Atomizdat, Moscow, 1973, 280. 31. Bregadze, V.G.; Nature of DNA interactions with cations: UV spectroscopic investigations and Marcus theory; Int. J. Quant. Chem., 1980, 17, 1213–1219. 32. Lavery, R.; Pullman, B.; The molecular electrostatic potential and steric accessibility of poly (dI.dC). Comparison with poly (dG.dC); Nucleic Acids Res., 1981, 9, 7041– 7052. 33. Lavery, R.; Cauchy, D.; De la luz, R.O. et al.; Molecular electrostatic potential of the BDNA helix. VII. Effect of screening by monovalent cations; Int. J. Quant. Chem., 1980, 7, 323–330. 34. Yakushevich, L.V.; DNA dynamics; Mol. Biol., 1989, 23, 652–662. 35. Frenkel, J.; Theorie der adsorption und verwandter erscheinungen; Z. Phys., 1924, 26, 117. 36. Frenkel, J.; Statistical Physics; M.-L. Acad. Sci. USRR, 1948, 760. 37. Slutsker, A.I.; Mikhailin, A.L.; Slutsker, I.A.; Microscopics of energy fluctuations of atoms in solids; Uspekhi Fizicheskikh Nauk, 1994, 164, 357–366. 38. Lineweaver, H.; Burk, D.; The determination of enzyme dissociation constants; J. Am. Chem. Soc., 1934, 56, 658–666. 39. Liliom K.; Orosz F.; Horvath L.; Ovadi J.; Quantitative evaluation of indirect ELISA. Effect of calmodulin antagonists on antibody binding to calmodulin; J. Immunol. Methods, 1991, 143, 119–125.

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40. Glaser, R.W.; Program for simulation of ELISA experiments and affinity; J. Immunol. Methods, 1993, 160, 129–133. 41. Rovati, G.E.; Rodbard, D.; Munson, P.J.; DESIGH: computerized optimization of experimental design for estimating K and B(max) in ligand binding experiments. I. Homologous and heterologous binding to one or two classes of sites; Anal. Biochem., 1988, 174, 636–649. 42. Scatchard, G.; The attraction of proteins for small molecules and ions; Ann. N.Y. Acad. Sci., 1949, 51, 660–672. 43. Klotz I.; Hunston,D.L.; Properties of graphical representations of multiple classes of binding sites; Biochem., 1971, 10, 3065–3069. 44. Gelagutashvili, E.S.; Rcheulishvili, A.N.; Mosulishvili, L.M.; Constants of Pb(II) ion binding to DNA at different ionic strengths; Biophysics, 2001, 46, 957–959. 45. Taylor, J.R.; An Introduction to Error Analysis; Mir, Moscow, 1985, 272. 46. Angelici, R.J.; Stability of coordination compounds. In: Inorganic Biochemistry, Vol. 1, G.L. Eichhorn, Ed., Mir, Moscow, 1978, Chap. 2, 89–132. 47. Scheiber, J.P.; Daune, M.; Interactions des ions metalliques avec le DNA. IV. Fixation de l’ion cuivrique sur le DNA; Biopolymers, 1969, 8, 139–152. 48. Frank-Kamenetski, M.D.; Anshelevich, V.V.; Lulashin, A.B.; Polyelectronic model of DNA; Uspekhi Fizicheskikh Nauk, 1987, 151, 595–618. 49. Gillespie, R.J.; Hargittai, I.; The VSEPR Model of Molecular Geometry; Allyn and Bacon, Boston, MA, 1991. 50. Shimoni-Livny, L.; Glusker, J.P.; Bock, Ch.W.; Lone pair functionality in divalent lead compounds; Inorg. Chem., 1998, 37, 1853–1867. 51. Gelagutashvili, E.; Studies of interaction between Ni(II), Zn(II) and Cr(III) ions and DNA from Spirulina platensis; Proc. Georgian Acad. Sci., Chem. Ser., 2005, 31, 69–73. 52. Gelagutashvili, E.; Belokobilsky, A.; Sigua, K.; Tarkashvili, C.; Nature of binding of Ni(II) and Zn(II) ions with DNA; Proc. Georgian Acad. Sci., Chem. Ser., 1999, 25, 107–111. 53. Nieboer, E.; Richardson, D.H.S.; The replacement of the non descript term ‘heavy metal’ by a biologically and chemically significant classification of metal ions; Environ. Pollution (Series B), 1980, 1, 3–26. 54. Rossetto, F.E.; Nieboer, E.; The interaction of metal ions with synthetic DNA: Induction of conformational and structural transitions; J. Inorg. Biochem., 1994, 54, 167–186. 55. Pauling, L.; General Chemistry; Mir, Moscow, 1974, 839. 56. Vologodskii, A.V.; Cozzarelli, N.R.; Conformational and thermodynamic properties of supercoiled DNA; Ann. Rev. Biophys. Biomol. Struc., 1994, 23, 609–643. 57. Jukhnevich, G.V.; Infrared Spectroscopy of Water; Nauka, Moscow, 1973, 207. 58. Granot, J.; Kearns, D.R.; Interaction of DNA with divalent metal ions. III. Extent of metal binding: experiment and theory; Biopolymers, 1982, 21, 219–232. 59. Sletten, E.; Lie, B.; Copper complex of guanosine-5′-monophosphate; Acta. Cryst., 1976, B32, 3301–3304. 60. Aoki, K.; General conclusion from solid state studies of nucleotide-metal ion complexes. In: Metal Ions in Biological Systems, A. Sigel and H. Sigel, Eds., Vol. 32, Marcel Dekker, New York, 1997, Chap. 4, 479–501. 61. Pohl, F.M.; Jovin, T.M.; Salt-induced co-operative conformational change of a synthetic DNA: equilibrium and kinetic studies with poly (dG-dC); J. Mol. Biol., 1972, 67, 375–396. 62. Pohl, F.M.; Polymorphism of a synthetic DNA in solution; Nature, 1976, 260, 365–366. 63. Sitko, J.C; Mateescu, E.M.; Hansma, H.G.; Sequence-dependent DNA condensation and the electrostatic zipper; Biophys. J., 2003, 84, 419–431. 64. Cantor, C.R.; Schimmel, P.R.; Biophysical Chemistry, Part 3, Mir, Moscow, 1985. 65. Bregadze, V.G.; Khutsishvili, I.G.; Structure of DNA complexes with the transition metals: UV difference spectra; Proc. Acad. Sci. Georgia, Biol. Ser., 1994, 20 (1–6), 272–280. 66. Freese, E.B.; Molecular Genetics, Part 1, J.H. Taylor, Ed., Academic Press, 1963, 135. 67. Drake, J.W.; Spontaneous mutations accumulating in bacteriophage T4 in the complete absence of DNA replication; Proc. Nat. Acad. Sci. USA, 1966, 55, 738–743.

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68. Danilov,V.I.; Quantum chemical Investigation of nucleic acid bases tautomer forms and problem of mutation; Biofizika, 1967, 12, 540–544. 69. Pullman, B; Pullman, A.; Le tautometrie des bases puriques et pyrimidiques et la theorie des mutations; Biochem. Biophys. Acta, 1962, 64, 403–405. 70. Pullman, B.; Pullman, A.; Electronic delocalization and biochemical evolution; Nature, 1963, 196, 1137. 71. Pullman, B.; Adenine complexes of thymine and uracil. In: Electronic Aspects of Biochemistry, B. Pullman, Ed., Academic Press, New York, 1964, 135. 72. Löwdin, P.O.; Effect of proton tunneling in DNA on genetic information and problems of mutations, aging and tumors; Biopolym. Symp., 1964, 1, 161–181. 73. Löwdin, P.O.; Some aspects of DNA replication; Incorporation errors and proton transfer. In: Electronic Aspects of Biochemistry, B. Pullman, Ed., Academic Press, 1964, 167. 74. Saenger, W.; Principles of Nucleic Acid Structure, Moscow, Mir; 1987, p. 584. 75. Bregadze, V.G.; Interpretation of ultra-violet difference spectrum of DNA complexes with metal ions of the first transition series; Biofizika, 1974, 19, 179–181. 76. Gralla, J.; Crothers, D.M.; Free energy of imperfect nucleic acid helixes, III. Small integral loops resulting from mismatches; J. Mol. Biol., 1973, 78, 301–319. 77. Dornberger, U.; Leijion, M.; Fritzsche, H.; High base pair opening rates in tracts of GC base pairs; J. Biol. Chem., 1999, 274, 6957–6962. 78. Bouvier, B.; Grubmuller, H.; A molecular dynamics study of slow base flipping in DNA using conformational flooding; Biophys. J., 2007, 93, 770–786. 79. Coman, D.; Russu, I.M.; A nuclear magnetic resonance investigation of the energetics of base pair opening pathways in DNA; Biophys. J., 2005, 89, 3285–3292. 80. Giudice, E.; Varnai, P.; Lavery, R.; Base pair opening within B-DNA: free energy pathways for GC and AT pairs from umbrella sampling simulations; Nucleic Acids Res., 2003, 31, 1434–1443. 81. Bhattacharya, P.K.; Cha, J.; Barton, J.K.; 1H NMR determination of base-pair lifetimes in oligonucleotides containing single base mismatches; Nucleic Acids Res., 2002, 30, 4740–4750. 82. Scott, R.A.; Scheraga H.A.; Method for calculating internal rotation barriers; J. Chem. Phys.; 1965, 42, 2209–2215.

3 Metal Ion Coordination in G-Quadruplexes Janez Plavec

3.1 Introduction DNA is able to adopt a variety of secondary structures other than the canonical BDNA which may potentially add considerable informational capacity to DNA.1–4 The initial concept of DNA being exclusively the holder of linearly arranged genetic information is being revisited to add alternative DNA conformations as important informational elements. Guanine nucleosides and nucleotides, as well as DNA (and RNA) sequences that include tracts of contiguous guanine residues form fourstranded structures known as G-quadruplexes (or tetraplexes or G4 structures). Several reviews on G-quadruplexes have been published recently5–12 including the monograph edited by Neidle and Balasubramanian.1 G-quadruplexes have been proposed to play roles in DNA transactions such as replication, transcription and recombination.13–15 These structures are adopted by G-rich nucleic acid sequences which have been found to be abundant throughout the genomes of many organisms.16–21 Potential G-quadruplex-forming regions are ample in telomers, oncogene promoter sequences and other biologically relevant regions of the genome. The fact that G-quadruplexes play roles in crucial biological processes makes them targets for therapeutic intervention.22–46 G-quadruplex-forming oligonucleotide sequences, which have the ability to selectively act as inhibitors of signal transduction or transcription via binding to particular targets, are known as aptamers.47–54 In addition to Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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their biological implications, the propensity for G-quadruplexes to form cationspecific supramolecular structures has opened the possibility for using guanine nucleoside and nucleotide analogues as well as G-rich DNA sequences for applications that range from environmental remediation to organic synthesis and nanotechnology.55–69 For example, molecular self-assembly of guanine-containing nucleosides has been shown to be useful for the construction of highly selective ionophores.70,71 G-rich sequences that fold into G-quadruplex structures can, of course, form Watson–Crick duplexes with their complementary C-rich strands. The formation of G-quadruplex or duplex structures within the cell will therefore depend on the relative stability of the G-quadruplex and the corresponding duplex.72–78 Furthermore, the C-rich strand might have the possibility of forming an intramolecular i-motif that likewise competes with duplex formation, which can also be influenced by cations or other changes in solution conditions (e.g. molecular crowding).79–83 The basic building block of a G-quadruplex is the G•G•G•G quartet, which is composed of four hydrogen-bonded guanine nucleotides in a horizontal planar arrangement (Figure 3.1). G-quartets are linked together by eight hydrogen bonds

Figure 3.1 The G-quartet: (a) four guanine residues that act both as acceptors and donors of H-bonds in Hoogsteen geometry form a cyclic G-quartet. R represents 2′-deoxyribose moiety; (b) side view of this planar building block of G-quadruplexes in cartoon presentation

Introduction

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in a Hoogsteen pairing geometry. The presence of cations is required for G-quartet formation. Cations reduce repulsions amongst guanine carbonyl oxygen atoms and contribute to enhanced base-base stacking interactions.5–8,10,84–88 The coordination of cations by the closely spaced carbonyl oxygen atoms of a G-quartet was postulated long before the first high-resolution structure of a G-quadruplex was determined. It is noteworthy that G-quartets interact with dehydrated cations via inner sphere coordination.89 It is therefore not surprising that formation, stability and structural details of G-quadruplexes are dependent on cation species and cation concentration. 3.1.1 Stoichiometry DNA G-quadruplexes can be formed with different numbers of oligonucleotide strands (i.e. 1, 2 or 4, Figure 3.2). In a tetramolecular structure four strands associate to form a G-quadruplex. Such quadruplexes are usually comprised of four strands in a parallel orientation (Figure 3.2a). Parallel quadruplexes have all guanine residues in an anti conformation. Association of two oligonucleotides with two G-tracts, which can be different, leads to a bimolecular structure with a so-called fold-back topology (Figure 3.2b). As a result, some of the G residues adopt the syn conformation and strands adopt different alternations of parallel and antiparallel orientations. The topological model in Figure 3.2b shows a G-quadruplex with two parallel and two antiparallel strands. In this structure each strand has one parallel and one antiparallel neighbour. An example of such an arrangement has been established experimentally by the structure of d[(G4T4G4)2] (see below). A monomolecular Gquadruplex can be adopted by sequences consisting of four G-tracts, separated by

Figure 3.2 (Plate 1) Variation in strand stoichiometry and topology of inter- and intramolecular G-quadruplex structures: (a) tetramolecular structure with all four parallel strands and all anti glycosidic torsion angles; (b) bimolecular and (c) monomolecular structures with antiparallel strands and alternating syn (in orange) and anti (in blue) orientations across glycosidic bonds (See colour plate section)

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three regions that can be any combination of residues, including G. An example of such an intramolecular structure with antiparallel strands and alternating syn and anti orientations across the glycosidic bonds is shown in Figure 3.2c. Quadruplexes are labelled as antiparallel when at least one of the four strands is antiparallel to the others. This type of topology is found in the majority of bimolecular and in many monomolecular G-quadruplex structures. Antiparallel folds exhibit both syn and anti guanine residues within individual G-quartets, which are arranged in a way that is in accordance with strand orientations. Recently, a formalism was proposed describing the interdependency of a set of structural descriptors as a geometric basis for folding of monomolecular Gquadruplex topologies.90 The formalism is a fundamental step towards the prediction of monomolecular G-quadruplex folding topologies from primary structures. In addition to establishing rules that will enable the prediction of folding topologies, the work of Webba da Silva90 suggested that there could be other folding topologies in addition to those that have been identified experimentally so far. 3.1.2 Loops G-quadruplex structures may also be classified according to the location of the loops that link the neighbouring G-tracts involved in G-quartets. The location and length of the loops in combination and in addition to the number of stacked G-quartets and the number and the pattern of the strand directionalities all contribute to a plurality of G-quadruplex structures. Edge-wise (or lateral) loops join adjacent Gstrands by spanning the edge of an outer G-quartet as shown schematically in Figure 3.3a. These loops are generally composed of two or more residues.91,92 This type of loop structure has been observed in two NMR solution-state structures of asymmetric G-quadruplexes of the d(TG4T2G4T) sequence93 and in the bimolecular Gquadruplex structure formed by the sequence d(G3CT4G3C).94 Two of the edge-wise loops can be located either on the same or on opposite sides of a G-quadruplex core, resulting in head-to-head or head-to-tail arrangements, respectively. Two distinct bimolecular G-quadruplexes were formed by d(G4T3G4).95 One of the structures exhibited a head-to-tail lateral loop dimer in which all adjacent strands were

Figure 3.3 Loop structures: (a) edge-wise loops connecting antiparallel strands on the same side of the G-quadruplex; (b) diagonal loops connecting opposite antiparallel strands; (c) external double-chain reversal loops connecting parallel strands

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antiparallel. The other was a head-to-head hairpin quadruplex with one adjacent strand parallel and the other antiparallel.95 The second type of loop joins opposite antiparallel G-strands by spanning the diagonal of the outer G-quartet (Figure 3.3b). Such loops usually consist of three or more residues. Diagonal loops were observed, for example, in the structure formed by the Oxytricha nova telomeric sequence d(G4T4G4).96–100 Adjacent parallel strands can be connected by a doublechain reversal loop that links the 3′-end of a given G-tract with the 5′-end of the other (Figure 3.3c). This loop arrangement gives a G-quadruplex a propeller type topology,101 with loops that can consist of a single or several residues.102–104 A doublechain reversal loop has been found both in crystal and solution-state structures for G-quadruplexes adopted by human telomeric DNA sequences, and more recently in a number of nontelomeric quadruplexes (see below). Residues in all three loop orientations have been shown to contribute to the stability of G-quadruplex structures though base-pairing and stacking over the neighbouring G-quartets.95,105–114 It is noteworthy that loops can adopt diverse structures beyond the basic classification mentioned above, which can be targeted by small molecule ligands.

3.2 Cation Coordination and Stability of G-Quadruplexes Early NMR studies of alkali metal ion specificity suggested that 5′-GMP formed self-ordered structures consisting of stacked G-quartets in the presence of K+, Na+ and Rb+, but not Li+ or Cs+.115 The first experimental evidence for direct coordination of dehydrated cations between G-quartets was provided by the observation of broadening and shielding of the 23Na resonance in solutions of 5′-GMP.116,117 K+ and Na+ ions are the most extensively characterized cations with respect to their ability to stabilize G-quadruplex structures.118–122 A list of other cations that have been shown to promote G-quadruplex formation includes the monovalent cations Rb+, Cs+,123,124 NH4+ 125,126 and Tl+100,127–129 as well as the divalent cations Sr2+,124,130–133 Ba2+,134,135 and Pb2+.125,136–138 In general, low divalent cation concentrations initially stabilize G-quadruplexes,139 while increasing concentrations eventually become destabilizing. Mn2+ and europium cations that stabilize G-quadruplex have been localized in the grooves.140,141 Studies of 8-bromoguanosine gel melting-transition temperatures indicated the following relative G-quartet stabilizing propensities of divalent cations: Sr2+ >> Ba2+ > Ca2+ > Mg2+. Davis and coworkers have shown that, in the presence of Pb2+, lipophilic guanine analogues can associate to form quadruplexlike structures in organic solvents.142 These results affirmed earlier studies that Pb2+ induces a more stable and compact structure compared to K+, which is evident from the comparison of cation-O6 bond lengths, O6-O6 diagonal distances and interquartet separation.136 Studies by Hardin and coworkers have shown that Mg2+ and Ca2+ ions stabilize the tetramolecular parallel-stranded G-quadruplexes formed by the sequences d(CGCG3GCG) and d(TATG3ATA) more than monovalent alkali cations.143,144 A CD spectroscopic study by Sugimoto and coworkers has shown that the bimolecular G-quadruplex d[(G4T4G4)2] formed in the presence of NaCl is actually destabilized by only 1 mM concentrations of Mg2+, Ca2+, Mn2+, Co2+, or

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Zn2+.76,145–147 Millimolar concentrations of Mg2+, Ca2+, Ba2+ and Sr2+ induce polymeric d(GGA)n repeat molecules to form quadruplexes.148 The studies of the melting temperature of gels formed by 8-bromoguanosine as a function of monovalent cation species showed that the relative ability to stabilize guanosine gels was K+ >> Rb+ > NH4+ > Na+ > Li+.149 The stabilization of G-quartet assemblies adopted by GMP follows the following order: K+ > NH4+ > Rb+ > Na+ > Cs+ > Li+.150 This general ordering has proven correct for G-quadruplex structures adopted by DNA oligonucleotides. Therefore, the melting temperature for a G-quadruplex is higher in the presence of K+ ions than in the presence of Na+ ions. The extent to which K+ increases the stability of a G-quadruplex over Na+ depends on the nucleotide sequence and can vary from as little as 2 °C to over 30 °C.107,124,151–155 The origins of these sequence-dependent differences are likely due to the K+ and Na+ forms of some G-quadruplexes being different in structure and/or in the number of coordinated cations. There is no simple explanation for the difference in melting temperatures for the Na+ and K+ forms for various G-quadruplexes. High-resolution X-ray and NMR structures have provided valuable insights into the effects of cation size and charge on G-quadruplex structure and on the location of ions within G-quadruplexes, all of which contribute to quadruplex stabilization.99,104,127,156–161 In addition, solution-state NMR studies have demonstrated that cations undergo dynamic exchange between coordination sites in the interior of a G-quadruplex and with bulk solution.162–165 The G-quadruplex formed by some G-rich sequences in the presence of K+ ions can be dramatically different in strand orientation and strand number from the Gquadruplex formed in the presence of Na+ ions. The size of a cation and its energy of (de)hydration both contribute to cation selectivity and the stability of a Gquadruplex. K+ is too large to be coordinated in the plane of the G-quartet, whereas a Na+ can fit in the centre of the G-quartet plane (see below). The resulting differences in K+ and Na+ coordination geometries and cation–guanine O6 distances contribute to a difference in the free energy provided by K+ versus Na+ coordination within a G-quadruplex. To a good approximation, hydration energies of monovalent ions are inversely proportional to their ionic radii.166 Thus, the free energy required to dehydrate K+ for coordination within a G-quadruplex is less than that required to dehydrate Na+. The net difference between the free energy of coordination within a G-quadruplex and the free energy of hydration ultimately determines cation selectivity by G-quadruplexes.6,153,167,168 A 1H NMR study by Hud and Feigon et al.168 on the bimolecular G-quadruplex adopted by d(G3T4G3) has shown that the preferred coordination of K+ over Na+ is actually driven by the greater energetic cost of Na+ versus K+ dehydration. The intrinsic free energy of Na+ coordination within d[(G3T4G3)2] is actually more favourable than K+ coordination.168 Subsequent computer simulations have provided additional support for the argument that cation dehydration is the dominant free energy that determines Na+ versus K+ selectivity by G-quartets.169–175 Recent quantitative analysis of the relative concentrations of the three dication forms of d[(G3T4G4)2] (i.e. 2 15NH4+, 15NH4+/K+ and 2 K+) at equilibrium, which are in slow exchange on the NMR timescale, showed that differences in standard Gibbs free energies are 5.7 kcal mol−1 between the di-15NH4+ and the 15NH4+/K+ forms, and 4.3 kcal mol−1 between the 15NH4+/K+ and the di-K+ forms.176 In comparison, the

Structure of Sequences Consisting of Human Telomere Repeats

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conversion of d[(G3T4G3)2] from its Na+ form to K+ form is associated with a net free energy change (DDG0) of 1.7 kcal mol−1.168 3.2.1 Molecular Switches In addition to their central role in stabilizing G-quadruplexes, cations also play an important role in determining G-quadruplex structure. A systematic study of G-rich oligonucleotides containing runs of G residues of various lengths and with varying numbers of intervening residues, revealed a cation-dependent pattern for the Gquadruplex folds.48,75,105,107,109,110,113,122,177–179 The possibility of cation-induced changes in G-quadruplex structures has formed the basis of the hypothesis that DNA Gquadruplexes might function as molecular switches.180,181 The interconversion of Grich DNA sequences between antiparallel and parallel G-quadruplexes could be controlled by Na+/K+ concentrations.182–186 Thomas and coworkers used Raman spectroscopy to generate a phase diagram of antiparallel and parallel G-quadruplex structures, as controlled by concentrations of Na+ and K+ ions.180 At low cation concentrations d[(T4G4)4] folds into an antiparallel diagonal-loop G-quadruplex. Higher cation concentrations and K+ ions lead to the formation of parallel structures.180 An entirely different ion-dependent conformational switch has been observed for d(G2AG2AG). At a moderate ionic strength of 150 mM NaCl, this oligonucleotide forms a four-stranded quadruplex.103 The bimolecular quadruplex comprises a unique A•(G•G•G•G)•A hexad motif that is stabilized by sheared GA base pairs, in addition to the G-quartet.103 At low ionic strength d(G2AG2AG) forms a twostranded V-shaped ‘arrowhead’ motif.187 The cation binding sites associated with the two folds of this oligonucleotide suggest a rational for the cation-dependent switch. A cation-dependent loop polymorphism has been observed in the bimolecular G-quadruplex adopted by d(G3CT4G3C) that contains G•C•G•C quartets, along with G-quartets.94,188 The conformation of thymines in the edge-wise loops is different between Na+ and K+ forms. The loop conformation is affected by the presence of a K+ binding site in the T4 loop, whereas there is no such binding site for Na+ ions. The loop conformation, in turn, is sensitive to the conformation of the outer G-quartet, which is substantially different in the two cation forms. A K+ ion is postulated to penetrate the G•C•G•C quartet at the expense of hydrogen bonding between the G-C base pairs.94,188 Some of the aspects of cation coordination within G-quadruplexes and their consequences that are mentioned and introduced in a rather general and limited way in the section above are further discussed in greater detail with specific examples in the rest of this contribution.

3.3 Structure of Sequences Consisting of Human Telomere Repeats Human telomeric DNA ends are composed of tandem TTAGGG repeats. It has been known for over a decade that G-quadruplexes formed from human telomeric

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repeat oligonucleotide sequences, e.g. d[(TTAGGG)4], exhibit different loop conformations depending upon whether Na+ or K+ ions are present.189 Studies on the structure and dynamics of the human telomeric G-quadruplex by single-molecule fluorescence resonance energy transfer (FRET) revealed two stable folded conformations in both Na+- and K+-containing buffers, with small differences between their enthalpies and entropies.74 Both folded conformations can be opened by the addition of a 21-mer complementary DNA oligonucleotide. The unfolding of both structures occurs at the same rate, which shows dependence on the monovalent cation present. Temperature-dependence studies in 100 mM KCl gave an apparent activation enthalpy and entropy of 6.4 kcal mol−1 and −52.3 cal mol−1 K−1, respectively, indicating that the unfolding is entropically driven and can occur easily. In contrast, in 100 mM NaCl the respective values are 14.9 kcal mol−1 and −23.0 cal mol−1 K−1, suggesting a more significant enthalpic barrier. These studies led to the conclusion that under near-physiological conditions, three structures coexist and can interconvert on a minute timescale.74,147,190–196 3.3.1 Single Repeat Sequences The oligonucleotide with a single repeat of the human telomere repeat sequence, d(TTAGGG), forms a tetramolecular parallel-stranded G-quadruplex at low K+ concentrations, which aggregates to form higher-order structures when the concentration of K+ is increased.197,198 3.3.2 Two-Repeat Sequences The X-ray structure of the two-repeat human telomeric sequence d(TAG3T2AG3T) formed in the presence of K+ ions exhibits a bimolecular structure with all strands in parallel orientation (PDB ID 1K8P).104 TTA residues form double-chain reversal (or propeller) loops. In addition, the end segments also participate in the formation of a T•A•T•A quartet, through pairing of the major groove edges of Watson–Crick A-T base pairs. K+ ions are coordinated between adjacent G-quartets.104 The same sequence has been shown to form both parallel and antiparallel G-quadruplex structures in the presence of K+ ions in solution.199 Both structures are bimolecular and comprise three stacked G-quartets. The two structures can coexist and interconvert in solution. They have different thermodynamic properties and different kinetics of folding and unfolding. d(TG4T2G4T), an analogous sequence with two G-for-A substitutions (originating from Tetrahymena), forms two asymmetric, bimolecular G-quadruplexes in solution that are involved in a similar equilibrium in the presence of Na+ ions.93 3.3.3 Four-Repeat Sequences The NMR structure of the monomolecular G-quadruplex formed by (almost) four repeats of the human telomere sequence, d[AGGG(TTAGGG)3], in the presence

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Figure 3.4 (Plate 2) Polymorphism of four repeats of human telomeric sequence: (a) intramolecular G-quadruplex formed by d[AG3(T2AG3)3] as determined by NMR spectroscopy in the presence of Na+ ions in solution;92 (b) the G-quadruplex fold with parallel GGG strands and double-chain reversal loops as determined by X-ray crystallography for the same sequence crystallized in the presence of K+ ions;104 (c) the topology of the four-repeat sequence in K+ ion containing solution (so called Hybrid-1 structure);204,205,207 (d) the fold of major form of the unmodified four-repeat sequence in K+ ion containing solution (so called Hybrid-2 structure).210 For clarity, only G-bases and their syn (in orange) and anti (in blue) orientations across glycosidic bonds are shown (See colour plate section)

of Na+ ions, was reported several years ago.92 This monomolecular G-quadruplex is stabilized by the three stacked G-quartets with anti-syn-syn-anti conformations around individual G-quartets (Figure 3.4a). A core of the three G-quartets is held together by strands in alternating orientations. Two of the TTA loops connect the edges of the outer G-quartet on the same side of the structure. The third loop is a diagonal type loop and is positioned on the other side of the G-quadruplex core. Such loop arrangements restrict access to the outer G-quartets by potential ligands. The 5′- and 3′-ends of the oligonucleotide are found at the same end of the Gquadruplex (Figure 3.4a). The crystal structure of the K+ form shows a dramatically different fold with all four GGG segments being parallel (Figure 3.4b).104 All guanine residues are in the anti conformation. All three TTA loops are in a double-chain reversal conformation and connect the top of one GGG strand with the bottom of the other. The loop residues are positioned alongside the grooves rather than at the ends of the Gquartet stack and thus give the topology a propeller shape. The double-chain reversal

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loops restrict access to the grooves, while the outer G-quartets represent large planar surface areas that were hoped to rationalize the selective binding of aromatic ligands. The 5′- and 3′-ends are at the opposite ends of the G-quadruplex (Figure 3.4b), which could support end-to-end stacking of successive G-quadruplexes in a larger DNA molecule. The crystal structure, in addition, revealed the position of K+ ions in the structure.104 In this 2.1 Å resolution crystal structure (PDB ID 1KF1) K+ ions are equidistant between the stacked G-quartets. The distances between K+ ions and each of the eight carbonyl oxygen atoms in the bipyramidal antiprismatic arrangement are 2.7 Å.104 Subsequently, several studies have demonstrated that an antiparallel Gquadruplex is formed by the human telomere repeat sequence in the presence of either Na+ or K+ ions in solution.74,192,200–202 In low salt conditions the human telomere DNA repeats tend to be either in disordered single-stranded states or parallel tetramolecular G-quadruplex structures.203 Determination of K+ ion structure in solution was elusive until recently when three groups reported the folding pattern of four repeats of human telomere sequence in the presence of K+ ions in solution.204–209 The main limitation for structure determinations was the existence of multiple conformers in equilibrium. The conformational equilibrium was driven into distinct predominant conformers that were amenable for structure determination by sequence modifications. Importantly, the modified oligonucleotide exhibited NMR spectral characteristics of the unmodified sequence. All these structures contain the (3 + 1) topology of the G-quadruplex core, in which three strands are oriented in one direction and the fourth is in the opposite direction (Figure 3.4c). The topology of this structure is different from the previously reported folds in the presence of Na+ ions in solution (Figure 3.4a) and K+ ions in the crystal (Figure 3.4b). This quadruplex contains three G-quartets with one exhibiting anti-syn-synsyn and two syn-anti-anti-anti conformations of guanine bases. One of the loops adopts a double-chain reversal structure, whereas the other two are edge-wise loops. In this fold, 5′- and 3′-ends of the sequence are located at opposite ends of the Gquadruplex core (Figure 3.4c). A related NMR study on a 26-nucleotide telomeric sequence d[(TTAGG)3TT] revealed that the wild-type human telomeric sequence adopts Hybrid 1 and Hybrid 2 conformations (Figures 3.4c and 3.4d) with a low energy barrier between them, which results in their coexistence in solution.210 Perusal of the topology of the Hybrid 2 structure shows the same (3+1) topology of the Gquadruplex core (Figure 3.4d). It consists of a single G-quartet with anti-syn-syn-syn and two with syn-anti-anti-anti conformations of guanine bases. The first two TTA loops closer to the 5′-end adopt an edge-wise structure, whereas the final loop adopts a double-chain reversal structure.

3.4 G-Quadruplexes Adopted by Promoter Regions Potential G-quadruplex-forming sequences are widespread in gene promoters. The analysis of bioinformatic data has shown that G-quadruplexes may be directly involved in gene regulation at the level of transcription. The promoter regions of

G-Quadruplexes Adopted by Promoter Regions 65

genes (1 kb upstream of the transcription start site) are significantly enriched in potential G-quadruplex motifs relative to the rest of the genome.21,211,212 Furthermore, the regions of the human genome that are both nuclease hypersensitive and within promoters show a remarkable 230-fold enrichment of G-quadruplex elements, compared to the rest of the genome.21 Sequences that have been studied for potential to form G-quadruplexes are c-myc,46,79,213–218 bcl2,219–222 VEGF,223 KRAS,224,225 RET,226 HIF-1a227 and c-kit.228–230 Structures adopted by these sequences, which are often composed of more than four G-tracts, are affected by their occurrence in the context of double-stranded DNA. The number of possible G-quadruplex structures is increased due to the fact that each of the G-tracts contains unequal number of guanines and that G-tracts are separated by different numbers of nucleotides. The c-myc gene and the respective protein are implicated in many cancers. The nuclease hypersensitivity element III1 controls up to 90% of total c-myc transcription. The guanine-rich sequence of this element contains six G-tracts, which result in multiple G-quadruplex folds that cannot be characterized individually by NMR. Early efforts in structure determination focused on the G-quadruplex that was formed by four of the six guanine tracts. Extensive search for suitable sequences afforded two sequences that exhibited sufficiently resolved NMR spectra, which corresponded to a single structure that enabled structure determination.231 One of the sequences named c-myc-2345 involved the second, third, fourth and fifth guanine tracts. The high-resolution structure of c-myc-2345 oligonucleotide in the presence of K+ ions has been determined by NMR (Figure 3.5a).231 This G-quadruplex represents an intramolecular propeller-type structure with all the strands in the parallel orientation. All guanines are in anti conformation. The structure consists of three G-quartets which are oriented in the same direction. All three loops are of doublechain reversal type. The structure resembles a parallel G-quadruplex with three double-chain reversal loops that was observed for the human telomere repeats in

Figure 3.5 (Plate 3) Representative structures of gene promoter regions: (a) G-quadruplex fold formed by guanine tracts of c-myc-2345 and c-myc-1245 sequences originating from the c-myc promoter, determined by NMR;231 (b) topology adopted by the c-myc-23456 sequence containing five of the six guanine tracts of the c-myc promoter region;232 (c) NMR structure of unprecedented intramolecular G-quadruplex formed by a G-rich sequence in the c-kit promoter in K+ solution233 (See colour plate section)

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the crystal (see below and Figure 3.4b). An interesting feature of this G-quadruplex is the occurrence of two single nucleotide double-chain reversal loops, which were first found in the case of the c-myc promoter G-quadruplex.44,221,231 The loop in the middle of the sequence of c-myc-2345 contains two nucleotides (GA) while the other two loops consist of a single T. The topology of a related c-myc-1245 sequence is similar to the structure shown schematically in Figure 3.5a. The difference between the structures of c-myc-1245 and c-myc-2345 is in the loop length. In the former, loops closer to the 5′- and 3′-ends of the molecule contain a single residue (A or T), while the central loop contains six residues (T5A). Interestingly, the G-quadruplex adopted by c-myc-2345 is more stable by 15 °C. The difference can be attributed to different loop length and consequently distinct structures. An NMR structure determination of c-myc-2345 mutant (two guanines replaced by thymines) adopted the topology shown in Figure 3.5a.216 It is important to note that the three structures mentioned above improved the insight into the structural details of c-myc promoter by correcting some earlier low-resolution assessments.46 The recent study focused on the c-myc sequence containing five of the six guanine tracts.232 The sequence designated as c-myc-23456 consisted of the second, third, fourth, fifth and sixth guanine tracts of the 27-mer c-myc NHE III1 element. The topology of the G-quadruplex adopted in the presence of K+ is shown in Figure 3.5b. The structure consists of three G-quartets. Interestingly, guanines involved in the G-quartets originate from each of the five tracts (underlined here: 5′-dTGA GGG TGG GGA GGG TGG GGA AGG-3′). The four strands are in parallel orientations. All guanines are in the anti conformation except for the 3′-end guanine, which is a snap-back syn nucleotide. The loops are in a double-chain reversal conformation, except for the final one, which is a diagonal loop and contributes to the stability of the structure (Figure 3.5b). A new G-quadruplex fold was shown to be adopted by the c-kit promoter sequence positioned upstream of the transcription start site (Figure 3.5c).233 The ckit promoter encodes for a tyrosine kinase receptor and is therefore involved in a regulation of signal transduction. c-kit kinase is targeted in clinical treatments of several cancers and by small molecule inhibitors that promote G-quadruplex formation.228,229,233–235 The c-kit sequence that folded into a single structure in the presence of K+ ions consists of four GGG tracts. Its structure exhibits an unusual feature where an isolated guanine is involved in the formation of a G-quartet core, despite the presence of four G-tracts (guanines involved in the G-quartets are underlined here: 5′-dAGG GAG GGC GCT GGG AGG AGG G-3′).233 Strands are in parallel orientation and all guanines are in the anti conformation. The structure contains four loops. Two of them are single-nucleotide loops in a double-chain reversal conformation. The two-residue linker connects two edges of the outer G-quartet, while the longer five-residue loop allows the final two guanines to be inserted into the G-quartets. The latter loop contributes to the stability of the structure through Watson–Crick A-T base pairs. The 5′ and 3′-ends are on the opposite sides of the G-quadruplex (Figure 3.5c), which is potentially of relevance in the context of longer DNA as it allows for continuation of the sequence. The use of single molecule fluorescence resonance energy transfer enabled detection of G-quadruplexes in the context of extended DNA duplex.234

Bimolecular G-Quadruplexes 67

3.5 Bimolecular G-Quadruplexes of Analogues of the Oxytricha Telomere Repeat The NMR structure of the G-quadruplex formed by d(G4T4G4) in the presence of NaCl was one of the first G-quadruplex structures determined at high resolution.96,97 DNA oligonucleotide d(G4T4G4), which contains 1.5 repeats of the Oxytricha telomere folds into a well-behaved bimolecular G-quadruplex. The topology, shown in Figure 3.6a, exhibits a symmetrical bimolecular G-quadruplex with four G-quartets and thymine loops at opposite ends of the G-quartets. The glycosidic torsion angles are syn-syn-anti-anti around each G-quartet. Guanines are alternately in syn and anti conformations along each strand. Adjacent strands are alternately in parallel and antiparallel orientations. The two T4 loops span across diagonals of the outer G-quartets.97,236 NMR solution studies showed that d(G4T4G4) adopts the same

Figure 3.6 (Plate 4) Topology of analogues of the Oxytricha nova telomere repeat sequence: (a) the structure of bimolecular d[(G4T4G4)2] quadruplex was one of the first high resolution structures of G-quadruplexes determined;96 (b) unique folding topology of d[(G3T4G4)2] with (3 + 1) strand orientations;244 (c) topology of d[(G4T4G3)2];243 (d) asymmetric structure of d[(G3T4G3)2]238 (See colour plate section)

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topology in the presence of K+ ions.98 A subsequent X-ray crystal structure of d[(G4T4G4)2] in the Na+ form, located in a cavity of the Oxytricha nova single-strand telomere-binding protein complex,158 revealed the same fold as the NMR structure. A recent X-ray study of the K+ form of d[(G4T4G4)2], which crystallizes in two crystal forms,99 also revealed identical folds to the NMR and the protein-associated structures. These results are likewise in accordance with the conformations deduced by Raman spectroscopy.237 The G-quadruplex formed by the related oligonucleotide d(G3T4G3) in the presence of K+ or Na+ ions has been determined independently by the groups of Feigon168,238 and Shafer239–241 by NMR in solution. d(G3T4G3) forms a bimolecular quadruplex (Figure 3.6d). Three G-quartets are formed from the GGG tracts. The conformation of guanine bases is syn-syn-anti-anti around each G-quartet. Adjacent strands are alternately in parallel and antiparallel orientations. Glycosidic conformations of the guanines are 5′-syn-syn-anti-(T4 loop)-syn-anti-anti in one strand and 5′-syn-anti-anti-(T4 loop)-syn-syn-anti in the other strand. The thymines in the loops span across the diagonal of the outer G-quartets. Unlike d[(G4T4G4)2], the bimolecular structure of d[(G3T4G3)2] is not symmetric, which results in separate signals for each monomer strand in NMR spectra. The unique feature of this topology is the stacking of the G-quartets, which includes both tail-to-tail and head-to-tail arrangements. The structures of d[(G4T4G4)2] (Figure 3.6a) and d[(G3T4G3)2] (Figure 3.6d) are closely related.97,98,185,240,242 However, the precise structure and dynamics of the T4 loops of both G-quadruplexes are cation dependent. A comparison of the Na+, K+ and 15NH4+ NMR structures of d[(G4T4G4)2] indicated that the coordination of Na+ ions within the planes of the outer G-quartets allows for the participation of the O2 carbonyl of the third loop thymine (T7) in the coordination of Na+ ions.98 In contrast, coordination of K+ or 15NH4+ ions between the planes of the G-quartets does not allow for coordination by the same loop thymine, which results in a different loop structure. When formed in the presence of K+ ions, there is also evidence of greater motion in the diagonal loops of the quadruplexes with respect to the Na+ structure.97,98,185,238,241 The sequence d(G4T4G3), which has one 3′ guanine missing from the sequence d(G4T4G4), has been shown to form a G-quadruplex with a significantly different structure and a greater cation-dependent polymorphism than d(G4T4G4).243 This sequence can be considered as an intermediate between d(G4T4G4) and d(G3T4G3). Our NMR study demonstrated that d(G4T4G3) forms a bimolecular G-quadruplex in the presence of Na+ ions. The topology of d[(G4T4G3)2], shown in Figure 3.6c, exhibits an asymmetric bimolecular fold-back structure consisting of three stacked G-quartets. The conformation of guanine bases is syn-syn-anti-anti around each Gquartet. The guanine bases of the two outer G-quartets exhibit a clockwise donor– acceptor hydrogen-bonding directionality, while those of the middle G-quartet exhibit anticlockwise directionality. The glycosidic torsion angle conformations of the guanine bases are 5′-syn–anti–syn–(anti-T4 loop)–anti–syn–anti in one strand and 5′-(syn)–anti–syn–anti–(T4 loop)–syn–anti–syn in the other strand. The two T4 loops both span diagonally across the outer G-quartets, but adopt different conformations. Each strand has neighbouring parallel and antiparallel strands. The two guanine residues not involved in G-quartet formation, G4 and G12 (i.e. the fourth

Coordination of Cations within d[(TG4T)4]

69

guanine base of one strand and the first guanine base of the other strand), adopt distinct conformations. G4 is stacked on top of an adjacent G-quartet and is part of the first loop. G12 is orientated away from the core of G-quartets and is stacked on the T7 base. The cation-dependent folding of the d[(G4T4G3)2] quadruplex structure is distinct from that observed for related sequences. While both d[(G4T4G4)2] and d[(G3T4G3)2] form bimolecular, diagonally looped G-quadruplex structures in the presence of Na+ and K+, we have observed this folding to be favoured for d[(G4T4G3)2] in the presence of Na+. The structure of d[(G4T4G3)2] exhibits a ‘slipped-loop’ element. The missing guanine residue at the 3′-end of d(G4T4G4), in terms of the primary structure, is compensated by slipping of the residues, which thus take its position within the 3D structure. Such a slipped strand apparently contributes to a thermodynamic stability of the structure. An NMR study on the folding of d(G3T4G4), a sequence with the 5′ terminal dG residue missing from the d(G4T4G4) sequence, established an unprecedented topology of a bimolecular G-quadruplex (Figure 3.6b).244 The structure consists of three G-quartets. The conformation of guanosine bases is anti-syn-syn-anti around one of the G-quartets, and syn-anti-anti-anti in the other two. Two guanine residues, G3 and G11, are involved in G-quartet formation. Three of the strands are in parallel, while one is in the antiparallel orientation. This type of topology resembles topological characteristics of (3 + 1) G-quadruplexes of the human telomere repeat sequences (see above). The outer G-quartets are spanned by diagonal as well as edgetype loops. Another unusual structural feature of d[(G3T4G4)2] is a leap between G19 and G20 over the middle G-quartet and chain reversal between G19 and G20 residues (Figure 3.6b). The examination of the influence of different monovalent cations on the folding of d(G3T4G4) showed that it forms a bimolecular G-quadruplex with the same general fold in the presence of K+, Na+ and ammonium ions.245

3.6 Coordination of Cations within d[(TG4T)4] Early studies on guanosine gels established a strong correlation between the melting temperature and the ionic radii of cations, which was an indication of site-specific ion binding by G-quartets.149 The strong interaction between cations and G-quartets originates from electrostatic interactions involving the guanine O6 oxygen atoms. On the other hand, electrostatic repulsions between cations within a G-quadruplex are substantial. Thus, the exact locations and coordination geometries of cations within a given G-quadruplex are the result of a balance between attractive interactions with carbonyl oxygen atoms and mutual cation repulsion. Cation coordination is not restricted to a particular geometry within a G-quadruplex (Figure 3.7). A series of stacked G-quartets produces a regular geometry, and potential cation coordination sites, with four O6 atoms within the plane of a G-quartet, or with eight O6 atoms between two stacked G-quartets. Ions such as K+ and 15NH4+ (ionic radii 1.33 Å and 1.43 Å, respectively) are too large to coordinate in the plane of a G-quartet, whereas Na+ (ionic radius 0.95 Å) is small enough to be coordinated within the plane of a G-quartet. Thus, as schematically illustrated in

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Figure 3.7 Sodium ion localization within G-quadruplexes as revealed by X-ray crystal structures of d[(TG4T)4] with eight strands forming two sets of four G-quartets157

Figure 3.7, multiple cation coordination geometries are possible and observed (see below). The determination of G-quadruplex crystal structures of d[(TG4T)4] has provided a detailed view of cation coordination that is unmatched by other DNA structures.157 The X-ray crystal structure of d[(TG4T)4] was refined to a resolution of 0.95 Å (PDB ID 352D). This crystal structure confirmed that d(TG4T) forms a tetramolecular parallel-stranded G-quadruplex in the presence of Na+ ions (Figure 3.8a).157 The asymmetric unit of the d[(TG4T)4] crystal consists of four Gquadruplexes, with two quadruplexes being coaxially stacked in a 5′ to 5′ orientation, such that a continuous stack of eight G-quartets is formed.157,246 Seven Na+ ions are coordinated along the axial channel formed by the O6 atoms of eight G-quartets (Figure 3.8a). Close inspection of the crystal structure reveals that the central Na+ in this channel is positioned equidistant between the planes of two stacked G-quartets. The coordination geometry of this particular Na+ is bipyrimidal, with eight equidistant guanine O6 atoms. Na+ ions located away from this central position are displaced incrementally further from this perfect symmetrical bipyrimidal coordination. At the ends of this channel, terminal Na+ ions are in fact located in the planes of the end G-quartets, coordinated by the carbonyl groups of only four guanine residues (Figure 3.8a). Displacement of the Na+ ions from the central eight-coordinate geometry to the in-plane four-coordinate geometry is believed to arise from electrostatic repulsion between adjacent Na+ ions. The mutual repulsion of Na+ ions along the central axis of the G-quadruplex has also been hypothesized to be the origin of an out of plane bending observed for the terminal G-quartets of the eight stacked quartets.157

Coordination of Cations within d[(TG4T)4]

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Figure 3.8 (Plate 5) Cation coordination within G-quadruplex structures adopted by d(TG4T) as revealed by X-ray crystallography: (a) Na+ ions (pink balls) within the 0.95 Å resolution structure (PDB ID 352D);157 (b) Na+ ions in the structure crystallized using lithium sulfate as the main precipitating agent (PDB ID 2O4F);246 (c) arrangement involving both Ca2+ (green balls) and Na+ (pink balls) cations crystallized in a mixed ion environment (PDB ID 2GW0);139 (d) Tl+ (red balls) and Na+ (pink ball) ions within the 2.2 Å resolution structure (PDB ID 1S45);247 (e) Tl+ (red balls) and Na+ (pink balls) ions within the 2.5 Å resolution structure (PDB ID 1S47);247 (f) Tl+ (red balls) and Na+ (pink balls) ions within the G-quadruplex exhibiting a T-quartet at the 3′-end of the strands (shown at the top of this structure) (PDB ID 1S47).247 Individual strands are shown in a wire frame model, while guanine bases are presented in stick representation to demonstrate the extent of out of plane bending. Cations are shown as coloured balls. Na+ with the smallest ionic radius can vary in coordination geometry from being within the plane of a G-quartet to being equidistant between two adjacent Gquartets. The larger ions are exclusively coordinated between two adjacent quartets (See colour plate section)

Analysis of details of the more recent crystal structure of d(TG4T) determined to 1.5 Å resolution (PDB ID 2O4F) shows that G-quartets within a bimolecular structure are not coplanar and are tilted with respect to the column axis towards the 5′-ends (Figure 3.8b).246 The crystallization of d(TG4T) from a solution containing both Li+ and Na+ ions showed only Na+ ions within the core of the quadruplex. A single bound Li+ ion has been observed at the surface of the bimolecular structure, but not within the G-quadruplex core.246 The high-resolution crystal structures of d(TG4T) also revealed the location of cations and water molecules located around the outside of the G-quadruplex.

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Crystals of d(TG4T) were also formed from a solution that contained both NaCl and CaCl2.157 As in many other nucleic acid crystals divalent Ca2+ ions were found at crystal lattice contact points between nucleic acid assemblies (i.e. bridging between backbone phosphates of two G-quadruplexes). Single water molecules were found to provide a fifth oxygen atom for coordination of the outermost Na+ ions that are coordinated in the planes of the terminal G-quartets. Additionally, water molecules were found along the phosphate backbone, in the four grooves, and as part of the hydration shell of Ca2+ ions. There was no evidence that Ca2+ ions substituted for Na+ ions at any sites within the G-quadruplex. Crystallization from a solution containing MgCl2 resulted in no bound divalent cations.246 An earlier study established a single Mg2+ ion located at the surface of the quadruplex.247 In the recent crystallographic study139 on d(TG4T) a new crystal form for the tetramolecular quadruplex with four parallel strands has been found (PDB ID 2GW0). In these crystals, formed in a mixed Ca2+ and Na+ ion environment, the channel along the central cavity of the quadruplex dimer is populated by cations in an asymmetric way (Figure 3.8c). One half has three Na+ ions, whereas the other half has three Ca2+ ions. The ion channel is capped by water molecules. Na+ ions are irregularly positioned with respect to the G-quartet planes. The outermost Na+ is almost coplanar with a G-quartet, whereas the inner one is almost equidistant from two G-quartets. A Ca2+ ion sits at the dimer interface (itself in plane with four other Ca2+ ions each in a groove). Two Ca2+ ions are approximately midway between G-quartets.139 d(TG4T) has been found to produce two distinct crystal forms when crystallized from solutions containing both Na+ and Tl+ ions.247 The quality and resolution of data (i.e. 2.2 and 2.5 Å) combined with the very different electron densities of Tl+ and Na+ ions allowed these two cations to be distinguished in the G-quadruplex structure. One crystal form contained two G-quadruplexes in the asymmetric unit (PDB ID 1S45), whereas the other form contained three G-quadruplexes (PDB ID 1S47). In all cases, G-quadruplexes are stacked in a head-to-head fashion with a 5′ to 5′ orientation, as found in the Na+ crystal structure.157 The main difference lies in the position of metal ions within the G-quadruplex and the low occupancy of innerG-quadruplex cation coordination sites by Tl+ ions (i.e. occupancy levels between 0.15 and 0.70). The Tl+ ions coordinated within the G-quadruplex are positioned between two neighbouring G-quartet planes (Figures 3.8d–e).247 This coordination geometry would be expected for Tl+ ions, as the ionic radius of Tl+ (1.44 Å) is slightly larger than that of K+ and certainly too large to be coordinated within the plane of a G-quartet. Low Tl+ ion occupancy has been attributed to the higher concentration of Na+ ions in the crystallization solution.247 The tilt of G-quartets towards the 5′-end junction has been observed in the Tl+-containing structure with one dimer (PDB ID 1S45), and with one dimer and one monomer (PDB ID 1S47).247 It has been appreciated for some time that Sr2+ ions can promote G-quadruplex formation.130 The fact that Sr2+ ions can stabilize G-quadruplexes to a similar degree as K+ seemed somewhat enigmatic. The ionic radius of Sr2+ (1.13 Å) is in between that of Na+ and K+, however the energy from electrostatic repulsions between Sr2+ ions within a G-quadruplex would be four times that of monovalent cations, if Sr2+ ions were coordinated with a similar spacing along the central axis of a

Coordination of Cations within d[(G4T4G4)2]

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G-quadruplex. The crystal structure of the RNA G-quadruplex formed by the sequence r(UG4U) in the presence of Sr2+ ions (PDB ID 1J8G), which was determined at ultra-high resolution (0.61 Å), seems to have provided an answer to this apparent enigma.160 Four strands of r(UG4U) form a parallel stranded quadruplex, similar to that described above for the analogous DNA sequence d(TG4T), but in the case of r(UG4U) the four strands within each quadruplex are symmetryrelated molecules. The tetramolecular G-quadruplexes of r(UG4U) stack with one another in opposite polarity (head-to-head or tail-to-tail) to form a pseudocontinuous column. Sr2+ ions are coordinated between every other pair of stacked G-quartets without any other cations coordinated at the intervening positions. The Sr2+ ions are associated with eight carbonyl oxygen atoms of adjacent G-quartets in a bipyramidal-antiprism geometry.160 Thus, it appears that the electrostatic repulsions between Sr2+ ions force these ions to leave vacant cation coordination sites within G-quadruplexes.

3.7 Coordination of Cations within d[(G4T4G4)2] 3.7.1 Crystal Structures The X-ray crystal structure at 1.86 Å resolution of the Oxytricha nova telomere end binding protein (OnTEBP) in complex with DNA and in the presence of NaCl has shown that d(G4T4G4) adopts a bimolecular G-quadruplex structure (PDB ID 1JB7)158 that is identical to the structure formed in the absence of the protein.99 Again, four guanine bases form four G-quartets, and four thymine residues form loops that span across the diagonal of the terminal G-quartets (Figure 3.6a). The structural features of the G-quadruplex in the protein–DNA complex are also very similar to those of the NMR structure,96,97 including the stacking of bases between G-quartets and the alternating syn-anti orientations across glycosidic bonds. The high-resolution X-ray diffraction data also reveals the positions of Na+ ions in the centre of the G-quadruplex.158 The two central Na+ ions are nearly coplanar within the central G-quartets and are in distorted, octahedral coordination environments (Figure 3.9a). The outer two Na+ ions are positioned above and below the planes of the outer G-quartets towards the T4 loops, and are coordinated by two O2 atoms of bases T5 and T7 in addition to the carbonyl oxygen atoms of the terminal G-quartet. Na+-thymine O2 coordination is consistent with the earlier proposal that changes observed by NMR in T nucleotide positions, when Na+ ions are exchanged for K+ or 15NH4+ ions, are driven by the loss of thymine O2 coordination.98 Electron density maps also revealed potential cation binding sites between the outside G-quartets and loops, which could not be unequivocally attributed to water molecules or Na+ ions.158 The bimolecular G-quadruplex formed by d(G4T4G4) in the presence of K+ ions has been crystallized in two different space groups and their structures have been refined to 2.0 and 1.49 Å resolution.99 One of these crystal structures (PDB ID 1JRN) has two G-quadruplexes per asymmetric unit, whereas the other crystal

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Figure 3.9 (Plate 6) Localization of cations within bimolecular G-quadruplexes: (a) Na+ ions (pink balls) within the d[(G4T4G4)2] quadruplex in crystals of the Oxytricha nova telomere end-binding protein with telomere DNA (PDB ID 1JB7);158 (b) K+ ions (grey balls) within d[(G4T4G4)]2 as revealed by X-ray crystal structure at 2.0 Å resolution (PDB ID 1JRN);99 (c) K+ ions (grey balls) within d[(G4T4G4)2] as revealed by X-ray crystal structure at 1.5 Å resolution (PDB ID 1JPQ);99 (d) five Tl+ ions (red balls) within d[(G4T4G4)2] as revealed by X-ray crystal structure at 1.55 Å resolution (PDB ID 2HBN);129 (e) ammonium ions within d[(G4T4G4)2] as established by NMR measurements in solution;162 (f) two ammonium ions within d[(G3T4G4)2] established by NMR studies in solution.176 Individual strands are shown in wire frame model, while guanine bases are presented in stick representation to demonstrate the extent of out of plane bending. Na+ ion can be localized within the plane of a G-quartet or between two adjacent G-quartets. The larger K+, Tl+ and 15NH4+ ions are exclusively coordinated between two adjacent quartets (See colour plate section)

structure (PDB ID 1JPQ) has only one G-quadruplex per asymmetric unit. All three G-quadruplex structures exhibit the same bimolecular diagonal loop folding topology. This fold is identical to the structure determined for the same DNA sequence ten years earlier by NMR in the presence of both Na+ and K+ ions (Figure 3.6a).96–98 The central two G-quartets in each G-quadruplex are approximately coplanar. One guanine base in each terminal G-quartet is slightly more tilted than the others and stacks effectively with the adjacent 3′-thymine base (Figures 3.9b and 3.9c). The average rise between adjacent G-quartets is consistently 3.3 Å.99 A linear row of five equidistant K+ ions lies along the helical axis within the central core of all three G-

Coordination of Cations within d[(G4T4G4)2]

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quadruplex crystal structures (Figure 3.9b). All K+ ions have full occupancy. The average K+–K+ distance is 3.38 Å. All three G-quadruplexes have three octahedrally coordinated K+ cations that are located equidistant between the planes of two adjacent G-quartets, with each K+ and eight O6 atoms forming a square antiprismatic arrangement. The outer K+ ions are located within the loops, where they also achieve octahedral coordination with the outer G-quartet O6 carbonyl oxygen atoms and the O2 atoms from the adjacent thymine bases, together with two oxygen atoms provided by water molecules. These potassium ions have only slightly increased mobilities (i.e. thermal factors) compared to the three central K+ ions.99 The crystal structure of the Tl+ form of the d[(G4T4G4)2] has been solved to 1.55 Å resolution (PDB ID 2HBN).129 This G-quadruplex contains five Tl+ ions (Figure 3.9d). The relative positions of these five cations are very similar to those found in the K+ crystal structure. The average distance between Tl+ ions is 3.6 Å. Three Tl+ ions are spread between adjacent G-quartet planes. They are coordinated by eight oxygens (one O6 from each of the surrounding guanines). Two Tl+ ions are coordinated in each of the two thymine loops (Figure 3.9d). They are coordinated by four guanine O6 atoms from the outer G-quartet plane and two thymine carbonyl groups. A water molecule was identified and located within the thymine loops and in close proximity to the loop-associated Tl+ ion.129

3.7.2 NMR Studies Ammonium ions have been shown to stabilize G-quartets to an extent that is similar to that observed for Na+.5,125 Hud and Feigon et al. used this fact to exploit 15NH4+ ion as a solution-state probe of monovalent cation coordination sites.162,248 These authors used 1H–1H and 1H–15N NMR correlation experiments to localize 15NH4+ ions within the bimolecular G-quadruplex d[(G4T4G4)2]. Dipole–dipole interactions (ROE) between the protons of the bound 15NH4+ ions and G-imino protons revealed that d[(G4T4G4)2] coordinates three 15NH4+ ions within its symmetric architecture (Figure 3.9e). In fact, only two distinct chemical shift environments were detected for bound 15NH4+ ions. One of them was assigned to the two symmetry-related outer sites, while the other was in the centre of the bimolecular G-quadruplex structure. The observed ROE cross-peak intensities also revealed that 15NH4+ ions in d[(G4T4G4)2] are positioned equidistant between stacked G-quartets (Figure 3.9e).162,176 NMR spectroscopy has also been used to monitor the competition between Na+ and 15NH4+ ions for coordination within G-quadruplexes at specific sites. For the G-quadruplex formed by d(G4T4G4) in the presence of 15NH4Cl, Na+ ions were shown to replace 15NH4+ ions at all three of the above mentioned outer- and innerbinding sites as the NaCl concentration is increased.163 Moreover, Na+ ions preferentially replace 15NH4+ ions coordinated at the inner site of d[(G4T4G4)2] quadruplexes.163 The 11-mer d(G3T4G4), a single base deletion of d(G4T4G4), has been shown by NMR spectroscopy to fold into an unusual asymmetric, bimolecular structure in the presence of 15NH4+, K+ and Na+ ions (Figure 3.6b).244 Multinuclear NMR studies

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have demonstrated that the three G-quartets of d[(G3T4G4)2] create two cation coordination sites (Figure 3.9f).176 Analysis of NOE interactions between the protons of the bound 15NH4+ ions and G-imino protons revealed that ammonium ions are localized between the pairs of adjacent G-quartets within d[(G3T4G4)2] (Figure 3.9f). Subsequent titration of KCl into a solution of d[(G3T4G4)2] folded in the presence of 15NH4+ ions revealed a mixed mono-K+/mono-15NH4+ form that represents an intermediate in the conversion of the di-15NH4+ form into the di-K+ form.176 Similarly, 15NH4+ ions were found to replace Na+ ions inside this G-quadruplex. The preference for 15NH4+ over Na+ ions for the two internal coordination sites is considerably smaller than the preference for K+ over 15NH4+ ions. Interestingly, the two coordination sites within d[(G3T4G4)2] differ to such a degree that 15NH4+ ions bound to the site that is closer to the lateral-type loop are always replaced first during titration by KCl. That is, the second binding site is not occupied by a K+ ion until a K+ ion already resides at the first binding site.176 Recently, 23Na NMR demonstrated the presence of mixed cation species (Na+/K+, Na+,Rb+ and Na+/Sr2+) of 5′-GMP associates in solution.249 A noticeable progress in localization of cations has been made by Wu and coworkers who demonstrated that tightly bound metal ions can be detected directly by 23Na, 39K and 87Rb NMR in solution.249,250 The observation of quadrupolar nuclei with characteristic rapid relaxation that has traditionally116,117,251 led to prohibitively broad lines has been made possible with the appearance of NMR spectrometers with high magnetic fields where separation of signals for channel and free cations is achievable. 23Na NMR study has confirmed that three Na+ cations reside inside the bimolecular G-quadruplex d[(G4T4G4)2].252 There was no evidence of Na+ cations in the thymine loop. Solid-state NMR spectroscopy has also been used to localize cations within G-quadruplexes.150,252–254 Distinct 23Na chemical shifts for Gquadruplexes formed by self-association of 5′-GMP were interpreted based on ab initio calculations on a G-quadruplex model consisting of four stacked Gquartets and three channel cations. In this model each cation is sandwiched between two adjacent G-quartets which are separated by 3.4 Å and twisted by 45 °. The 23Na chemical shielding calculations with Na+ ion located within the G-quartet plane predicted 23Na chemical shift (d + 6 ppm), which does not agree with the experimental data (d − 17 ppm).249 It is noteworthy, however, that the localization of Na+ ions solely between the neighbouring G-quartet planes is not fully supported by the available crystal structures discussed above and shown in Figures 3.8 and 3.9. Tl+ has been shown to nicely substitute for K+ in the promotion of Gquadruplex formation.100,127,128 205Tl is a spin-½ nucleus with relatively high receptivity, in contrast to other monovalent cations. Its potential advantage could be a very large chemical shift range and thus high sensitivity to the chemical environment of a metal cation. As expected, the resonances of 205Tl+ ions associated with G-quartets were clearly resolved from the resonance of 205Tl+ ions in bulk solution. Strobel and coworkers have reported the NMR spectra for 205Tl+ ions in a solution containing the tetramolecular G-quadruplex d[(T2G4T2)4].127 The 205Tl NMR spectrum of this G-quadruplex shows a cluster of three peaks.127 The observation of three distinct 205 Tl peaks of approximately equal areas is consistent with a tetramer in which three Tl+ ions are localized between the layers of the four G-quartets. The ability of Tl+

Coordination of Cations within d[(G4T4G4)2]

77

ions to compete with Na+ ions for coordination within the G-quadruplex d[(G4T4G4)2] has been verified by solution-state 1H NMR.128 Differences in resonance peak line widths for the three Tl+ resonances suggests that ions move from one cation binding site to the other. The central peak was narrower, which was attributed to the slower exchange of the Tl+ ion at the middle coordination site. The outside Tl+ ions exchange with bulk solution more freely. The lifetime of Tl+ ions bound within the Gquadruplex was estimated to be at least 3 ms.127 One of the first NMR studies to determine the number of cations bound within a G-quadruplex in the solution state was performed using the sequence d(G3T4G3).168 In the presence of either NaCl or KCl, d(G3T4G3) forms a bimolecular fold-back structure (Figure 3.6d).238,240 1H NMR spectroscopy was used to follow the competition between Na+ and K+ ions for coordination sites within the G-quadruplex.168 Changes in 1H NMR spectra during these titration experiments indicated a gradual transition of d[(G3T4G3)2] from the G-quadruplex structure observed in the presence of only NaCl to the G-quadruplex structure observed in presence of only KCl. Although the Na+ and K+ forms of d[(G3T4G3)2] have the same molecular folds, there are relatively small structural differences between the two forms that are manifested by changes in 1H chemical shifts. No separate or additional 1H resonances were observed in samples of d[(G3T4G3)2] that contained various mixtures of NaCl and KCl, which demonstrated that the exchange rate of Na+ and K+ forms of this quadruplex at 25 °C is fast on the NMR timescale (ca. <10 ms). 1H chemical shift changes observed over the course of KCl titration into a sample of d[(G3T4G3)2] were fit perfectly by a model in which two Na+ ions are replaced by two K+ ions within the G-quadruplex. Furthermore, quantitative analysis of chemical shift changes during the same titration experiments168 gave important insights regarding the thermodynamic basis for the preferential coordination of K+ over Na+ by G-quadruplexes (see below). d[G4(T4G4)3] has been folded into a monomolecular G-quadruplex in the presence of 15NH4+ ions (Figure 3.2c).164,255 Topology of the folded structure is equivalent to the solution-state structure in the presence of Na+ ions determined earlier in the laboratories of Patel256 and Feigon.257 The 15NH4+ ion binding sites within the d[G4(T4G4)3] G-quadruplex were identified and assigned with the use of 2D 1H–15N HSQC and NOESY spectra.164 The 3D structure of this monomolecular Gquadruplex consists of four G-quartets with alternating parallel and antiparallel strands (Figure 3.2c).256,257 The conformation of guanines along the strands alternates between syn and anti. One of the T4 loops spans diagonally across the outer Gquartet, while the two T4 loops are in an edge-wise conformation. Three 15NH4+ ion binding sites were established within the interior of d[G4(T4G4)3].164 There was no indication of 15NH4+ ions residing at the loop binding sites. The volumes of HSQC cross-peaks corresponding to the three binding sites were not the same. The outer site (named O1), which is closer to the outer G-quartet spanned by the diagonal loop showed the highest occupancy. The inner binding site, I, and the other outer site, O2, demonstrated comparable occupancies at 130 mM 15NH4Cl.164 Quantitative analysis afforded the equilibrium binding constants of 93, 31 and 46 M−1 for O1, I and O2 binding sites.255 The binding site I which is in the core of d[G4(T4G4)3] exhibits the lowest affinity for 15NH4+ ions. The binding site O1 exhibits a three times higher

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equilibrium binding constant, which suggests that the terminal 3′ and 5′ guanines of d[G4(T4G4)3] within the outside G-quartet facilitate 15NH4+ ion binding to this site.

3.8 Cation Movement within G-Quadruplexes NMR spectroscopy has been used extensively to measure the dynamics of cations coordinated within G-quadruplexes. By analysing resonance linewidths in NMR spectra of the quadrupolar nuclei 23Na and 39K, Braunlin and coworkers were able to estimate the rotation times and resident bound lifetimes of dehydrated cations within G-quadruplexes d[(T2G4T)4] and d[(G4T4G4)2].258,259 Cations within these two G-quadruplexes have rotation times that are on the order of nanoseconds. The rotational immobilization exhibited by coordinated cations within G-quadruplexes is in distinct contrast to the high rotational mobility of cations associated with the surface of a G-quadruplex. Na+ ions coordinated within the tetramolecular quadruplex d[(T2G4T)4] were shown to exchange more rapidly compared to Na+ ions coordinated within the bimolecular quadruplex d[(G4T4G4)2].259 The lifetime of Na+ ions coordinated within d[(G4T4G4)2] was estimated to be 250 µs at 10 °C.259 The residence lifetimes determined for Na+ ions tightly bound within DNA quadruplexes are in distinct contrast to the kinetics of G-quartet base opening, which can be as slow as days to weeks.110,152,236,260 Thus, cations must move between bulk solution and the cation coordination sites within a G-quadruplex by passing along the axial channel of the quadruplex, rather than by moving through transient side openings in G-quartets. Wu et al. have observed similar 23Na NMR spectra for d[(TG4T)4] at two other magnetic fields, 9.4 and 11.7 T.250 At 9.4 T, a chemical shift separation between the channel and surface (or bulk) Na+ cations of 17 ppm corresponds to a frequency separation of approximately 1800 Hz. This chemical shift difference suggests that the residence time for Na+ cations inside the G-quadruplex channel of d[(TG4T)4] is ~0.6 ms at 25 °C.259 The fact that a separate 23Na NMR signal is observed for the channel Na+ ions with Mn+ (K+, Rb+ and Sr2+) as an immediate neighbour suggests that the movement of the added Mn+ ions in and out of the G-quadruplex channel must be slow on the 23Na NMR timescale.249 The smallest signal separation observed for the different channel Na+ ions is 0.6 ppm (corresponding to 95 Hz at 14.1 T) observed in the Na+/K+ case. This indicates that the average residence time of K+ ions inside the channel must be much longer than ~2 ms.249 This is consistent with the recent results that the residence time of 15NH4+ ions in a d[(G4T4G4)2] quadruplex channel containing mixed Na+/15NH4+ ions is 36 ms,163 and that the residence time of Tl+ ions in the same G-quadruplex is approximately 100 ms.100,127 NMR studies utilizing 15NH4+ ions as a probe for localizing cation binding sites and for monitoring the movement of cations between different coordination sites revealed that 15NH4+ ions move along the central axis of the bimolecular quadruplex d[(G4T4G4)2] and the solution, reminiscent of an ion channel.161–165,248 The residence time of 15NH4+ ions at the inner site is 250 ms at 10 °C. 15NH4+ ions were shown to move between the three inner binding sites, and from the outer binding sites into

Cation Movement within G-Quadruplexes 79

solution. The relatively large size of 15NH4+ ions in comparison to Na+ ions is likely to hinder their movement through the quadruplex and provides a rationale for its longer residence time compared with Na+ ions. The significant difference in the residence times of 15NH4+ and Na+ ions bound inside the bimolecular G-quadruplex d[(G4T4G4)2] revealed that 15NH4+ ion movement from the inner to the outer binding sites of d[(G4T4G4)2] is accelerated by ca. 7.5 times in the presence of Na+ ions.163 The residence time of 15NH4+ ions within the inner coordination site is reduced from 270 ms in the presence of 15NH4+ ions alone, to 36 ms in the presence of both 15NH4+ and Na+ ions at 25 °C.163 The partial occupancy by Na+ ions is strongly correlated with the acceleration of 15NH4+ ion movement through the G-quadruplex, whereas bulk ion concentration does not exert a noticeable influence on the rate of 15NH4+ ion movement from the inner to the outer coordination sites. Even a low concentration (i.e. 5 mM) of Na+ ions decreases the residence time of 15NH4+ ions bound at the inner coordination site to the minimum value of 36 ms. Further increases in Na+ concentrations, to the point that one or two coordination sites of d[(G4T4G4)2] are completely occupied by Na+ ions, does not lead to further increase in the rate of 15 NH4+ ion movement. Thus, the residence time of 36 ms for 15NH4+ ions bound at the inner coordination site provides a good estimate for the limiting rate for its movement from the inner to an outer binding site. The much slower rate of 15NH4+ ion movement observed when all three coordination sites of d[(G4T4G4)2] are occupied by 15NH4+ ions is apparently due to the added restriction that their movement to an adjacent site within the G-quadruplex requires a concerted movement of another 15NH4+ ions out of the adjacent site (i.e. double 15NH4+ ion movement).163 Vacant coordination sites are likely to exist, at least transiently, during 15NH4+ ion movement. However, the lifetime of these vacancies must be short compared with the lifetime of occupied sites. An NMR study on 15NH4+ ion movement within d[(G3T4G4)2] demonstrated the absence of 15NH4+ ion movement between the two binding sites along the central ion cavity of the G-quadruplex structure in solution.165 15NH4+ ions do move between the two binding sites, U and L, and bulk solution. 15NH4+ ion movements from the binding sites U and L into the bulk solution are characterized by lifetimes of 139 ms and 1.7 s at 298 K, respectively. The 12 times faster movement from binding site U demonstrates that 15NH4+ ion movement is controlled by the structure of T4 loop residues, which through diagonal- versus edge-type orientations impose distinct steric restraints for cations to leave or enter the G-quadruplex. Activation energy for the UB process is 15.8 kcal mol−1.165 The d[G4(T4G4)3] quadruplex exhibits four G-quartets with three 15NH4+ ionbinding sites (termed O1, I and O2).164 The analysis of 2D NMR data has demonstrated unambiguously that there is no unidirectional 15NH4+ ion movement through the central cavity of the d[G4(T4G4)3] quadruplex.164 15NH4+ ions move back and forth between the binding sites within the G-quadruplex and exchange with ions in bulk solution. 15NH4+ ion movement is controlled by the thermodynamic preferences of individual binding sites, steric restraints of the G-quartets for 15NH4+ ion passage and diagonal versus edge-type arrangement of the T4 loops. The cross-peak volumes were observed for mixing times as long as 3 s, which was due to slow 15N longitudinal relaxation and slow 15NH4+ ion movement within the d[G4(T4G4)3] quadruplex. The

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movement of 15NH4+ ions from the interior of the G-quadruplex to bulk solution is faster than exchange within the G-quadruplex. Residence lifetimes of 20 and 14 s, respectively, were established for the movement of 15NH4+ ions from the two outer binding sites into the inner binding site at 10 °C. A temperature increase to 40 °C led to shortening of residence lifetimes to 1.5 and 2.0 s for the O1I and O2I processes, respectively. The temperature-dependent rate constants afforded activation energies of 15.4 and 12.1 kcal mol−1 for O1I and O2I processes, respectively. 15NH4+ ion movement between the inside of the G-quadruplex and bulk solution is faster from the O2 than the O1 binding site (Figure 3.2c, binding site O2 is the one closer to the two edge-wise loops). It seems that structural details of the d[G4(T4G4)3] Gquadruplex define the stiffness of individual G-quartets that intimately affect 15NH4+ ion movement. The stiffness of G-quartets and steric hindrance imposed by thymine residues in the loops contribute to the five-fold difference in the exchange rate constants through the outer G-quartets.

3.9 Summary Our insights into the possible biological roles of G-quadruplexes and into the origins of their topological sensitivity to cation species and concentration have benefited from recent studies on structure, stability and interactions of cations with constituent G-quartets. However, it is not currently possible to delineate general rules governing the folding of G-rich DNA sequences, and more specifically the role of cations in this process. Different G-rich sequences adopt distinct quadruplex topologies and, in addition, a given sequence can also fold into various different conformations, which can coexist. Human telomere sequences represent one of the best illustrations of the structural diversity and complexity of G-quadruplex structures. The change of metal ion species present can induce structural changes, which can be significant for some DNA sequences. A simple change in cation (salt) concentration can result in large changes in the structure and stability of a G-quadruplex. NMR and X-ray crystallography contribute important insights into the occupancy of binding sites by cations and kinetics of ion movement, which are intrinsically correlated with the structural details, dynamic fluctuations and local flexibility of the DNA structure. In general, X-ray crystal structures demonstrate that cations such as K+ and Tl+ are coordinated exclusively between the adjacent G-quartets because they are too large to be coordinated within the plane of a G-quartet. In contrast, smaller Na+ ions can be coordinated within a G-quartet with three distinct ligand geometries: (i) bipyrimidal coordination sites directly between two G-quartet layers, (ii) octahedral coordination sites where the ion is coplanar with the G bases and (iii) intermediate, less symmetric positions. Inter-Na+ distances in the parallelstranded tetramolecular structure adopted by d(TG4T) and the bimolecular structure adopted by d(G4T4G4) are greater than the average distance between stacked G-quartets. Studies on the relationship between G-quadruplex stability and cation species have primarily considered ionic radius as the most important factor. However, the fact that Na+ and Ca2+ have almost the same ionic radii illustrates that there are

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other factors, including the energy of dehydration and/or coordination number, which must be considered. Thermodynamic stability data and cation competition experiments consistently show that K+ ions have an optimal balance between favourable coordination within a G-quadruplex and not too unfavourable free energy of dehydration. At the other end of this spectrum are Cs+ and Li+ ions. It appears that these cations are not favourable for G-quadruplex stabilization because Cs+ is too large for optimal coordination by G-quartets, whereas the free energy required for Li+ dehydration is much higher than for the other alkali metal ions.

Acknowledgements Financial support from Slovenian Research Agency (ARRS) and the Ministry of Higher Education, Science and Technology of the Republic of Slovenia (Grant Nos. P1-0242 and J1-0986) is gratefully acknowledged.

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187. Kettani, A.; Bouaziz, S.; Skripkin, E.; Majumdar, A.; Wang, W.M.; Jones, R.A., et al.; Interlocked mismatch-aligned arrowhead DNA motifs; Structure, 1999, 7, 803–815. 188. Bouaziz, S.; Kettani, A.; Patel, D.J.; A K cation-induced conformational switch within a loop spanning segment of a DNA quadruplex containing G-G-G-C repeats; J. Mol. Biol., 1998, 282, 637–652. 189. Balagurumoorthy, P.; Brahmachari, S.K.; Structure and stability of human telomeric sequence; J. Biol. Chem., 1994, 269, 21858–21869. 190. Lee, J.Y.; Okumus, B.; Kim, D.S.; Ha, T.J.; Extreme conformational diversity in human telomeric DNA; Proc. Natl. Acad. Sci. USA, 2005, 102, 18938–18943. 191. Qi, J.Y.; Shafer, R.H.; Human telomere quadruplex: Refolding and selection of individual conformers via RNA/DNA chimeric editing; Biochemistry, 2007, 46, 7599–7606. 192. Qi, J.Y.; Shafer, R.H.; Covalent ligation studies on the human telomere quadruplex; Nucleic Acids Res., 2005, 33, 3185–3192. 193. Yu, H.Q.; Miyoshi, D.; Sugimoto, N.; Characterization of structure and stability of long telomeric DNA G-quadruplexes; J. Am. Chem. Soc., 2006, 128, 15461–15468. 194. Miyoshi, D.; Karimata, H.; Sugimoto, N.; Drastic effect of a single base difference between human and Tetrahymena telomere sequences on their structures under molecular crowding conditions; Angew. Chem. Int. Ed. Engl., 2005, 44, 3740–3744. 195. Pedroso, I.A.; Duarte, L.F.; Yanez, G.; Burkewitz, K.; Fletcher, T.M.; Sequence specificity of inter- and intramolecular G-quadruplex formation by human telomeric DNA; Biopolymers, 2007, 87, 74–84. 196. Antonacci, C.; Chaires, J.B.; Sheardy, R.D.; Biophysical characterization of the human telomeric (TTAGGG)4 repeat in a potassium solution; Biochemistry, 2007, 46, 4654–4660. 197. Wang, Y.; Patel, D.J.; Guanine residues in d(T2AG3) and d(T2G4) form parallel-stranded potassium cation stabilized G-quadruplexes with anti glycosidic torsion angles in solution; Biochemistry, 1992, 31, 8112–8119. 198. Kato, Y.; Ohyama, T.; Mita, H.; Yamamoto, Y.; Dynamics and thermodynamics of dimerization of parallel G-quadruplexed DNA formed from d(TTAG(n)) (n = 3–5); J. Am. Chem. Soc., 2005, 127, 9980–9981. 199. Phan, A.T.; Patel, D.J.; Two-repeat human telomeric d(TAGGGTTAGGGT) sequence forms interconverting parallel and antiparallel G-quadruplexes in solution: Distinct topologies, thermodynamic properties, and folding/unfolding kinetics; J. Am. Chem. Soc., 2003, 125, 15021–15027. 200. Redon, S.; Bombard, S.; Elizondo-Riojas, M.A.; Chottard, J.C.; Platinum cross-linking of adenines and guanines on the quadruplex structures of the AG3(T2AG3)3 and (T2AG3)4 human telomere sequences in Na+ and K+ solutions; Nucleic Acids Res., 2003, 31, 1605–1613. 201. He, Y.J.; Neumann, R.D.; Panyutin, I.G.; Intramolecular quadruplex conformation of human telomeric DNA assessed with I-125-radioprobing; Nucleic Acids Res., 2004, 32, 5359–5367. 202. Li, J.; Correia, J.J.; Wang, L.; Trent, J.O.; Chaires, J.B.; Not so crystal clear: the structure of the human telomere G-quadruplex in solution differs from that present in a crystal; Nucleic Acids Res., 2005, 33, 4649–4659. 203. Rujan, I.N.; Meleney, J.C.; Bolton, P.H.; Vertebrate telomere repeat DNAs favor external loop propeller quadruplex structures in the presence of high concentrations of potassium; Nucleic Acids Res., 2005, 33, 2022–2031. 204. Ambrus, A.; Chen, D.; Dai, J.X.; Bialis, T.; Jones, R.A.; Yang, D.Z.; Human telomeric sequence forms a hybrid-type intramolecular G-quadruplex structure with mixed parallel/antiparallel strands in potassium solution; Nucleic Acids Res., 2006, 34, 2723–2735. 205. Luu, K.N.; Phan, A.T.; Kuryavyi, V.; Lacroix, L.; Patel, D.J.; Structure of the human telomere in K+ solution: An intramolecular (3 + 1) G-quadruplex scaffold; J. Am. Chem. Soc., 2006, 128, 9963–9970.

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Metal Ion Coordination in G-Quadruplexes VEGF gene by the presence of potassium and G-quadruplex-interactive agents; Nucleic Acids Res., 2005, 33, 6070–6080. Cogoi, S.; Xodo, L.E.; G-quadruplex formation within the promoter of the KRAS proto-oncogene and its effect on transcription; Nucleic Acids Res., 2006, 34, 2536– 2549. Cogoi, S.; Quadrifoglio, F.; Xodo, L.E.; G-rich oligonucleotide inhibits the binding of a nuclear protein to the Ki-ras promoter and strongly reduces cell growth in human carcinoma pancreatic cells; Biochemistry, 2004, 43, 2512–2523. Guo, K.; Pourpak, A.; Beetz-Rogers, K.; Gokhale, V.; Sun, D.; Hurley, L.H.; Formation of pseudosymmetrical G-quadruplex and i-motif structures in the proximal promoter region of the RET oncogene; J. Am. Chem. Soc., 2007, 129, 10220–10228. De Armond, R.; Wood, S.; Sun, D.Y.; Hurley, L.H.; Ebbinghaus, S.W.; Evidence for the presence of a guanine quadruplex forming region within a polypurine tract of the hypoxia inducible factor 1 alpha promoter; Biochemistry, 2005, 44, 16341–16350. Fernando, H.; Reszka, A.P.; Huppert, J.; Ladame, S.; Rankin, S.; Venkitaraman, A.R., et al.; A conserved quadruplex motif located in a transcription activation site of the human c-kit oncogene; Biochemistry, 2006, 45, 7854–7860. Rankin, S.; Reszka, A.P.; Huppert, J.; Zloh, M.; Parkinson, G.N.; Todd, A.K., et al.; Putative DNA quadruplex formation within the human c-kit oncogene; J. Am. Chem. Soc., 2005, 127, 10584–10589. Todd, A.K.; Haider, S.M.; Parkinson, G.N.; Neidle, S.; Sequence occurrence and structural uniqueness of a G-quadruplex in the human c-kit promoter; Nucleic Acids Res., 2007, 35, 5799–5808. Phan, A.T.; Modi, Y.S.; Patel, D.J.; Propeller-type parallel-stranded G-quadruplexes in the human c-myc promoter; J. Am. Chem. Soc., 2004, 126, 8710–8716. Phan, A.T.; Kuryavyi, V.; Gaw, H.Y.; Patel, D.J.; Small-molecule interaction with a fiveguanine-tract G-quadruplex structure from human MYC promoter; Nat. Chem. Biol., 2005, 1, 167–173. Phan, A.T.; Kuryavyi, V.; Burge, S.; Neidle, S.; Patel, D.J.; Structure of an unprecedented G-quadruplex scaffold in the human c-kit promoter; J. Am. Chem. Soc., 2007, 129, 4386–4392. Shirude, P.S.; Okumus, B.; Ying, L.M.; Ha, T.; Balasubramanian, S.; Single-molecule conformational analysis of G-quadruplex formation in the promoter DNA duplex of the proto-oncogene c-kit; J. Am. Chem. Soc., 2007, 129, 7484–7485. Bejugam, M.; Sewitz, S.; Shirude, P.S.; Rodriguez, R.; Shahid, R.; Balasubramanian, S.; Trisubstituted isoalloxazines as a new class of G-quadruplex binding ligands: Small molecule regulation of c-kit oncogene expression; J. Am. Chem. Soc., 2007, 129, 12926–12927. Smith, F.W.; Feigon, J.; Strand Orientation in the DNA Quadruplex Formed from the Oxytricha Telomere Repeat Oligonucleotide d(G4T4G4) in Solution; Biochemistry, 1993, 32, 8682–8692. Krafft, C.; Benevides, J.M.; Thomas, G.J.; Secondary structure polymorphism in Oxytricha nova telomeric DNA; Nucleic Acids Res., 2002, 30, 3981–3991. Smith, F.W.; Lau, F.W.; Feigon, J.; d(G3T4G3) forms an asymmetric diagonally looped dimeric quadruplex with guanosine 5′-syn-syn-anti and 5′-syn-anti-anti N-glycosidic conformations; Proc. Natl. Acad. Sci. USA, 1994, 91, 10546–10550. Scaria, P.V.; Shire, S.J.; Shafer, R.H.; Quadruplex structure of d(G3T4G3) stabilized by K+ or Na+ is an asymmetric hairpin dimer; Proc. Natl. Acad. Sci. USA, 1992, 89, 10336–10340. Keniry, M.A.; Strahan, G.D.; Owen, E.A.; Shafer, R.H.; Solution structure of the Na+ form of the dimeric quadruplex [d(G3T4G3)]2; Eur. J. Biochem., 1995, 233, 631– 643. Strahan, G.D.; Shafer, R.H.; Keniry, M.A.; Structural properties of the [d(G3T4G3)]2 quadruplex: evidence for sequential syn-syn deoxyguanosines; Nucleic Acids Res., 1994, 22, 5447–5455.

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4 Supramolecular Chemistry of Metal–Nucleobase Complexes P. J. Sanz Miguel, P. Amo-Ochoa, O. Castillo, A. Houlton and F. Zamora*

4.1 Introduction Studies of the interaction of metal ions with nucleic acids are central to bioinorganic chemistry and this area continues to be actively researched. As well as providing valuable insight into the mode of action of numerous metal-containing drugs (e.g. cis-platin), understanding these interactions has provided opportunities for exploiting the nucleobases as highly versatile ligands. The basic coordination chemistry of these purine and pyrimidine derivatives has been summarized in several reviews.1 During the last four decades, more than 600 metal–nucleobase complexes have been prepared and crystallographically characterized. The number of crystal structures reported to date for metal ions with nucleobases as ligands has increased dramatically in recent years reflecting the great interest in these materials (Figure 4.1). The structural characteristics of nucleobases allow extension beyond basic coordination chemistry towards larger, more complex, molecular architectures based on the formation of supramolecules. Two broad types of supramolecules are of particular interest in the development of new discrete and infinite structures; firstly those formed exclusively through coordinate bond formation involving the nucleobases and, secondly, supramolecules which self-assemble through complementary Hbonding of nucleobases acting as ligands to metal centres.2

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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Figure 4.1 Historical evolution of the metal–nucleobase structures deposited on the CSD database. Each colour represents a year of that decade

The aim of this chapter is to provide an overview of supramolecular aspects of transition metal–nucleobase chemistry. This is not an exhaustive review, but rather summarizes the structural possibilities that have been described through the combination of these two types of components: metal ions and nucleobases. As well as this, a brief overview of the biological, chemical and physical properties of these compounds is given. The formation of a particular supramolecular architecture requires the selection of suitable building blocks. Supramolecular systems require their components to have appropriate spacing and configuration relative to each other, and ideally a library of related structural units is required. The structural subunits for the construction of coordination supramolecules are the metal ions, or complexes, and organic ligands. Architectures based on these systems have been the subject of numerous investigations due to their potential for applications based on their useful properties, such as molecule and ion hosts, optical properties, porous materials and catalytic activity.3 Geometrical aspects have a critical role in determining the final architecture of the supramolecular entity. Specifically, in the case of metal ions (or complex fragments) and nucleobases two different types of building blocks are used for constructing finite or infinite molecular assemblies; the most typical syntheses proceed in a single step by self-assembly, however, in a few cases multistep synthetic approaches have been shown more convenient, i.e. nonlabile metal fragments. 4.1.1 Nucleobases as Building Blocks Nucleobases are ideally suited as building blocks for supramolecular coordination chemistry due to their versatility as ligands. The most relevant factors in this respect are: (i) the large number of donor sites available for metal binding, (ii) the Hbonding donor/acceptor sites and (iii) the potential for p–p stacking, analogous to

Introduction

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Figure 4.2 Essential purine and pyrimidine nucleobases

Figure 4.3 Theoretical binding angles of the nucleobases towards metal ions

that observed in helical duplex DNA. Together these bond-making characteristics provide a rich basis from which to synthesise supramolecular architectures. Nucleobases are divided in two groups: purines and pyrimidines (Figure 4.2). The larger purine moiety provides an increased number of donor sites (e.g. endocyclic N1, N3, N7, N9). Different metal ion coordination patterns establish a range of possible angles between the metal–ligand bond vectors. Figure 4.3 exhibits possible coordination angles for purines and pyrimidines, respectively. Deviation from expected values of these angles between coordinate bond vectors increases the structural possibilities.

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Figure 4.4 Finite and infinite supramolecules can be achieved by combination of ditopic linear and angular building blocks, or by combining different linear building blocks

4.1.2 Metal ions and Complexes as Building Blocks Coordination of transition metal ions or complex fragments to nucleobases permits the assembly of either discrete geometric entities or polymeric structures. The selection of metal ions, or complex fragments, is based on several factors. Most important is the ability to display the appropriate coordination geometry for the desired architecture of the final supramolecule. These geometries may be considered broadly as either linear or angular, as exemplified by the use of cis- and trans-amine complexes of platinum, with angular (90 °) and linear (180 °) geometries. Figure 4.4 gives a summary of infinite architectures of coordination supramolecules obtained by combination of linear and angular subunits.

4.2 Discrete Architectures Many metal fragments offer a variety of specific angles for construction as a consequence of their known coordination geometries. However, metal fragments with bond angles at 90 or 180 ° are the most used with nucleobases, to form discrete, or closed, structures. Formation of finite architectures require the self-assembly of either angular or a combination of angular and linear building blocks. Figure 4.5 provides a summary of discrete supramolecules that have been formed using nucleobases and metal ions as building blocks.

Discrete Architectures

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Figure 4.5 Feasible ways to form finite architectures (triangles, squares, hexagons) using ditopic ligands (O) and metal ions (M)

4.2.1 Triangles Purines A molecular triangle has recently been synthesized by Lippert et al.3a in a selfassembly process of the metallo fragment trans-[(NH3)2Pt(Pu-N7)(H2O)]2+ (Pu = 9methylpurine), previously formed by reacting trans-[(NH3)2Pt(OH)Cl] with 9-methylpurine followed by halide abstraction with AgNO3. 1H NMR reveals that although trans-[(NH3)2Pt(Pu-N7)(H2O)]2+ is formed initially, this subsequently

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Figure 4.6 Schematic representation of the trans-[{(NH3)2Pt(µ-N1-Pu-N7)}3]6+ molecular triangle and the trans-[{(NH3)2Pt(µ-N1-Pu-N7)}4]8+ molecular square

condenses at room temperature to form short chains that evolve, after five days, to give two different closed complexes: the trans-[{(NH3)2Pt(m-N1-Pu-N7)}3]6+ molecular triangle and the trans-[{(NH3)2Pt(m-N1-Pu-N7)}4]8+ molecular square, in a 1 to 0.6 ratio (Figure 4.6). It is noteworthy that the N1–Pt–N7 angles in the trimer deviate from the ideal 180 ° to 168.8–170.8 ° and the angles between the Pt atoms coordinated to N1 and N7 sites of the purine ligands deviate significantly from 90 ° to 67.8–70.8 °. These substantial geometric deviations allow the formation of the unexpected triangular arrangement. The ratio of products can be reversed to 1 : 2.5 (triangle:square) by addition of Na2SO4 and this also has the effect of enhancing the reaction rate. In the X-ray structures of the compounds, the purine ligands and the platinum(II) atoms are essentially coplanar and the ammine ligands are oriented perpendicular to the former plane. In addition, both complexes provide molecular cavities capable of anion recognition with particular affinity for sulfate. Few examples of organometallic complexes with nucleobases are known. Cyclic trimers resembling hollow truncated cones have been synthesized by the reaction of trimethylplatinum(IV) acetate with 9-methyladenine (9-MeAH) in a 1 : 1 ratio4. The complex presents three PtMe3 moieties bridged by deprotonated 9-methyladenine ligands which are bound to one platinum via N6 and N7 and via N1 to another platinum (m-κN1:κ2N6,N7) (Figure 4.7). Thus, the platinum centres are octahedrally coordinated by three methyl ligands and three nitrogen donor atoms of the adenines. Crystals from different solvents show slight differences in the structural details of the trimeric unit and in crystal packing. The reaction between the metal fragments (h6-arene)Ru(II) (arene = C6H6, pcymene = CH3C6H4CH(CH3)2) and (h5-arene)M (M = Rh(III) and Ir(III); arene = pentamethylcyclopentadienyl) with hypoxanthine and adenine derivatives provides attractive water-soluble cyclic trimeric (Figure 4.7) and tetrameric complexes.5–8 The ability of these types of cyclic supramolecular complexes as receptors have been studied, and binding of biological molecules has been observed. In particular, the Rh(III) cyclic trimers with adenine derivatives recognize amino acids by a variety

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Figure 4.7 Representation of the main features of the triangular supramolecules formed with Pt(II) and Ru(II) and 9-Methyladenine or 9-Methylhypoxanthine

3+

Figure 4.8 Essential skeleton of the triangle [( PMe3 )2Pt (1-MeC ) ( H − )]3

of noncovalent processes involving p–p, hydrophobic and H-bonding interactions. Formation of these trimeric structures is achieved by coordination of the purine to metal ions as: (i) m-[h2(N7,O6):h1(N1)] with Rh(III) and 9-ethylhypoxanthine; (ii) m-[h2(N7,N6):h1(N1)] with Ir(III), Rh(III) and 9-EtA, 9-MeA, respectively. It has been found that when the 9 position of adenine is not blocked, formation of a similar cyclic tetramer complex take places via m-[h2(N7,N6):h1(N9)]. Pyrimidines Preliminary 1H, 13C and 195Pt NMR spectra of the products obtained by the reaction of 1-methylcytosine (1-MeC) with cis-[(PMe3)2Pt(m-OH)]2(X)2 ( X = NO3−, ClO4− ) in water, suggested that single mononuclear complexes, cis-[(PMe3)2Pt(1-MeC−)](X), were formed. However, X-ray crystallographic analyses identified the products as the trinuclear cis-[(PMe3)2Pt(1-MeC−)]3(X)3 ( X = NO3−, ClO4− ) , formed by deprotonation of the exocyclic amino group of the 1-MeC. Natural abundance 15N NMR results are in agreement with the structural data obtained in the solid state.9 The initial reaction products are dinuclear complexes, cis-[(PMe3)2Pt(1-MeC−)]2(X)2, which rearrange to a thermodynamically more stable trinuclear species in water or DMSO at 80 °C. The driving force for the oligomerization reaction has to be related to a more favourable metal–ligand interaction in the trinuclear species, likely due to the lower intramolecular repulsion of the cis-(PMe3)2Pt moieties. The X-ray structure of the perchlorate complex shows three cis-(PMe3)2Pt units symmetrically bridged by cytosinate anions through the N3 and N4 atoms (Figure 4.8). The

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platinum atoms lie at the corners of a virtually equilateral triangle (mean Pt···Pt separation of 5.31 Å with mean Pt–Pt–Pt angle of 60 °). The nucleobases are strictly planar, their centroids are placed equally far from the centre of the triangle and the plane of the centroids is practically parallel to that of platinum atoms.9 4.2.2 Squares As for other metallomacrocycles based on metal fragments and donor ligands, several examples with nucleobases as building blocks have been reported, in which triangles and squares are in equilibrium in solution. Frequently, these equilibria can be modulated by addition of neutral or anionic species. An example has already been presented in the previous section3a. In all these cases the squares represent the most thermodynamically stable species compared to the triangle. Purine Houlton and coworkers have developed the coordination chemistry of chelatetethered nucleobase derivatives. These contain a chelating group, such as ethylenediamine, tethered to the N9-position of purines through a short alkyl chain (as shown in Figure 4.9). The presence of this flexible metal binding group has given rise to some unique tetrameric structures upon reaction with metal ions. For example, reaction of CdSO4 and en-Et-G (G-Et-en = ethylenediamine-N9-ethylguanine) yields the tetramer-containing salt, [Cd4(G-Et-en)4(H2O)4(SO4)2], shown in Figure 4.9.10 In this compound the metal–nucleobase bonding involves N7 alone. The ethylenediamine group acts to bridge adjacent metal ions forming the discrete tetrameric assembly. Additional ligands, water and sulfate anions act to produce columns throughout the solid-state structure. The tetrameric assembly contains four Cd(II) centres lying in the same plane, effectively forming the corners of a square. The dimensions of the square are: Cd1···Cd2 edge 8.729 Å, Cd1···Cd2A edge 7.970 Å, Cd1···Cd1 diagonal 11.451 Å, Cd2···Cd2 diagonal 12.176 Å. The Cd–Cd–Cd angles

Figure 4.9 Schematic representation of the guanine-based square in [Cd4(N7-G-Et-en)4 (H2O)4(SO4)2]

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Figure 4.10 Schematic illustration of the guanine-based square [Pd4(N3,N7-G-prop-en)4]8+

are 86.5 and 93.5 °. This structure is also described as a one-dimensional polymer with SO42− acting as bridging ligands (see Section 4.3.1). Tetrameric molecular boxes are generally formed through cis-ligand orientation at the metal centre or with the N1,N7-bridging mode of the purines that also present a 90 ° angle. However, in [Cd4(G-Et-en)4(H2O)4(SO4)2] N7 and the chelating tether effectively providing orthogonal bond vectors10 (Figure 4.9). Another example of a tetrameric complex was isolated from the reaction of Pd(II) ions with en-Prop-G (G-Prop-en = ethylenediamine-N9-propylguanine11). In this case a distorted molecular box structure is observed, which features coordination of both N7 and N3 of the guanine moiety, as well as the ethylenediamine group (Figure 4.10). The complex may be considered as an octacationic metallomacrocycle. The Pd···Pd edge distances exhibit a narrow range between 7.237 and 7.269 Å. The diagonal distances are Pd1···Pd3 9.871 Å and Pd2···Pd4 10.062 Å and the four palladium ions lie in a plane. As in the Cd(II) tetramer, the guaninyl residues act to form approximately linear bridges, though in this case coordination also involves N3. This is a rare metal binding site for guanine with a very limited number of examples reported.11 A unique aspect of these guanine-based tetramers is that, in contrast to the natural tetrads (telomeres), which are assembled through hydrogen bonding and metal ion stabilization, these structures are formed through metal–ligand bonding alone. Since the sites on the nucleobase involved in coordination are N7 or N7 + N3, this leaves the Watson–Crick face available for base pairing (Figure 4.11). This feature presents numerous opportunities for preparing supramolecular metal–DNA hybrid structures assembled via complementary H-bonding interactions. It also raises the interesting question of how these compounds might interact with natural DNA strands.11 Pyrimidines The self-assembly of pyrimidine ligands as angular building blocks (120 °) to metal fragments with binding angles at 90 ° gives rise to metallocalix[4]arene formation. These are metal analogues to the classical calix[4]arenes with a typical geometry

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Figure 4.11 Scheme illustrating the potential for base pairing between the Cd-based squares in [Cd4(N7-G-Et-en)4(H2O)4(SO4)2] and complementary cytosines

Figure 4.12 Scheme of 1,3-alternate and cone conformations of metallocalix[4]arenes

that resembles an open box. Several metallocalixarenes with pyrimidine ligands show that different configurations coexist in solution: cone, 1,3-alternate and intermediate conformers (Figure 4.12). The principal interest in calixarenes is based on their ability to function as hosts for small molecules or ions, as well as their metal binding properties, either in the cavity or at the periphery. By the application of

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Figure 4.13 Representation of the tetranuclear cation {[Pt(en)]4(UH-N1,N3)4]}4+ and the molecular boxes formed by coordination of metal ions

cis-configured metal complexes, e.g. Pt(II) and Pd(II), as metal building blocks and pyrimidine nucleobases as bridging ligands, it is possible to obtain new multinuclear complexes. The most common architectures of metallocalixarenes with nucleobases reported so far have been cyclic tri- and hexanuclear supramolecules. The combination of the 90 ° binding angle provided by the [Pt(en)]2+ fragments and the angle of 120 ° of the N1,N3 coordination of uracil allows the formation of tetranuclear metallocycles, which may be considered as special metallocalix[4]arenes. These complexes are able to interact with cations, via the exocyclic donating oxygen atoms located at the 2- and 4-positions, and to act as receptors of organic anions in the central cavity. The X-ray analysis shows four [Pt(en)]2+ entities, which occupy the corners of a square (Pt···Pt 5.86 Å), bridged by four uracil rings such that each pyrimidine molecule is bound through N1 to a platinum atom and through N3 to a second one (Figure 4.13). The coordination geometry about the Pt(II) is normal square planar with slight angle deviations. The addition of cis-[(NH3)2Pt]2+, [Pt(en)]2+, [Pd(en)]2+, Ni(II), Cu(II) and Ag(I) to [(en)Pt(UH-N1,N3)]4(NO3)4 yields octanuclear complexes. In all the octanuclear compounds the basic structure of the open box is retained, but instead of a single [Pt(en)]2+ residue, two metal units, PdII(en) or MII, represent the corners of the boxes (Figure 4.13).12–13 4.2.3 Larger Assemblies Purines The reaction between cis-[(PMe3)2Pt(NO3)2] and 9-MeGH (9-methylguanine), in aqueous solution, produced the first example of a cyclic hexanuclear platinum– guanine derivative, cis-[(PMe3)2Pt(9-MeG)]6(NO3)6 in which six cis-(PMe3)2Pt units are bridged by deprotonated guanine ligands through their N1 and N7 atoms (Figure

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Figure 4.14 Structural representation of the [(PMe3)2Pt(9-MeG))]6(NO3)6 molecular hexagon

4.14).14 The platinum atoms lie in two parallel planes separated by 0.34 Å, and these planes make an angle of 50 ° with the plane of the nucleobases. The distance between two adjacent metal atoms is 6.5 Å, and the longest Pt···Pt distance is 12.9 Å. Hydrogen bonding plays an important role in stabilizing the hexanuclear cation and the crystal lattice, which is destroyed in absence of the mother liquor. Although it is not comprised of hexagons, there is another example of a complex based on a windmill-shaped hexanuclear copper(II) cluster {(H2O)2Cu2(m3-(A)4 [Cu(oda)(H2O)]4}·6H2O (oda = oxydiacetato), in which a dinuclear ‘core’ [CuII2( µ-N 3, N 9-A )4( H 2O)2 ] is connected to four peripheral Cu(oda)(H2O) units (Figure 4.15).15 The dimeric fragment is centrosymmetric and is made of two CuII atoms bridged by four m-N3,N9-adeninate anions in a windmill-shaped arrangement ′ . This structural subunit was previously reported with a Cu···Cu distance of 2.94 Å for some dimeric compounds.16 Pyrimidines Rauterkus et al., have synthesized the first pentanuclear metallomacrocycles with pyrimidines (thymine (T) and uracil (U)).17 The reaction between 2,2′-dipyridylketonedichloridoplatinum(II) with an excess of uracil or thymine affords three pentanuclear platinum(II) macrocyclic compounds. Three different Pt(II) entities are found in these structures in which the uracil or thymine ligands are coordinated to platinum through the endocyclic nitrogens (N1,N3) and also by means of the exocylic oxygens (O2 and/or O4) (Figure 4.16). The compounds have the ability to capture different anions (nitrate, perchlorate, tetrafluoroborate and hexafluoro-

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Figure 4.15 Hexanuclear {(H2O)2Cu2(µ3-(Ade)4[Cu(oda)(H2O)]4}·6H2O complex with windmill topology

Figure 4.16 Schematic representation of the pentanuclear metallomacrocycles formed between the 2,2′-dipyridylketoneplatinum(II) entity and thymine or uracil

phosphate), via hydrogen bond formation between the exocyclic oxygen atoms of the coordinated nucleobases and the anions. The chelate ligands 2,2′-dipyridiylketone and 2,2′-dipyridiylketone hydrate provide further functional groups, supplementary to those of the DNA bases, for the formation of a dense hydrogen-bond network. These characteristics provide new potential applications in anion recognition and complexation. A molecular hexagon was obtained using the trans-[(Pt(CH3NH2)2(1MeC2−)2](NO3)2 (1-MeC2− = dianion of 1-methylcytosine), fixing a head–head arrangement by metal binding (with HgII) to the deprotonated exocyclic amino groups. Electrophilic attack of excess Hg(II) salts at the C5 positions of the two cytosine ligands leads to the cyclic [{Pt(CH3NH2)2(C5H3N3O)2Hg3(OH)(NO3)}2] (NO3)4. The basic form of this centrosymmetric cation is a compressed hexagon

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Figure 4.17 Schematic representation of the [{Pt(CH3NH2)2(1-MeC)2Hg3(OH)(NO3)}2] (NO3)4 molecular hexagon.

(sides 7 and 5.5 Å) with four HgII and two PtII atoms at the edges and four nucleobases and two OH groups representing the corners (Figure 4.17). All metal ions and OH groups are almost coplanar. A special structural characteristic of this hexagon is the arrangement of the six HgII ions. The interactions between the Lewis acidic Hg centres and nucleophiles could lead to specific substrate binding and catalysis.18

4.3 Infinite Architectures As with discrete supramolecules, polymers may also adopt different architectures (see Figure 4.4) depending on the choice of metal and the nucleobase binding sites. The design and construction of polymeric compounds are of great interest due to their ability to act as functional materials, such as nonlinear optical (NLO), electrical switches or materials which possess interesting magnetic or catalytic properties. Three-dimensional polymers may also show applications in molecular storage (gases, anionic interchange etc.) (Figure 4.18). The formation of polymers based on biologically relevant ligands such as nucleobases, may provide additional applications in biological fields. For instance, some of these structures could be used as models of specific forms of DNA (e.g. M-DNA, G4). Nucleobase-derived coordination polymers may be classified in three different groups: (i) the nucleobases act as unique bridging ligands between the metal fragments, (ii) the nucleobases and a second ligand type link the metal centres or (iii) the nucleobases are terminal ligands coordinated to the metals, which are connected through other bridging ligands. A general observation is the relatively small number of polymeric structures based on bridging nucleobases alone, in spite of the great number of available coordination positions. A feasible explanation for this fact could be the use of nucleobases in their neutral form, which imposes the presence of counterions to balance the charge of the metal ions. At the same time, the anions compete with the nucleobases for binding to the metal centres. This could, in principle, be overcome by the deprotonation of the bases, improving their coordination ability and avoiding the need for counterbalancing anions.

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Figure 4.18 Possible applications of infinite nucleobase–metal architectures

Figure 4.19 Scheme of the structure of the 1D coordination polymer obtained by coordination of Ag(I) entities to pyrimidinato ligands

4.3.1 1D Coordination Polymers Nucleobases as Bridging Ligands With 1-methyluracil in its anionic form, the Aoki group prepared a silver-based polymeric compound, coordinated through the carbonyl groups and the deprotonated N3 of the pyrimidinato ligands (Figure 4.19).19 Beauchamp and coworkers reported another similar example: the reaction of Ag(I) with 1-methylthymine leads to the formation of the 1D coordination polymer via deprotonation of the nucleobase, which acts as a bridging ligand between two silver ions. Two different Ag(I) ions are observed in the polymeric chain, one with linear coordination involving two N3 donor atoms and one tetrahedral Ag(I) centre coordinated to two O2 and two O4 atoms from four thymines.20 With uracilato as ligand, the Lippert group isolated a heteronuclear Pt/Ag polymer. The PtAg2 entities are linked through O2 and O4 of the uracilato ligands,

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Figure 4.20 Structure of the 1D coordination polymer [(1-MeU-N3,O2,O4)2Ag2Pt]n

Figure 4.21 Schematic representation of the linear chain of {[Ag(1-MeC)](NO3)}n

as shown in Figure 4.20.21 This complex represents one of the few examples of infinite architecture based on nucleobases with two different metals. The reaction of AgNO3 with 1-methylcytosine (1-MeC) results in the polymeric {[Ag(1-MeC)](NO3)}n complex. This contains centrosymmetric dimers in which the 1-MeC ligands are bridged by two Ag(I) coordinated to O2 and N3 (Figure 4.21). The nitrate ion is also coordinated to the silver ion. The dimeric units [Ag(1MeC)(NO3)]2 form columns where the nucleobases are stacked. Based on this structure, new models for the binding of Ag(I) to regions of high G–C content in duplex DNA were proposed.22 Another interesting example, obtained by Lippert using platinated nucleobases as building blocks for supramolecular entities, is the product from the reaction of trans-[(NH3)2Pt(9-MeA-N7)(9-MeHx-N7)](ClO4)2 with AgNO3. This yields a 1D helical coordination polymer trans-[{(NH3)2Pt(9-MeA-N7)(9-MeHxN7)}Ag(NO3)(H2O)] (ClO4)(NO3) (Figure 4.22). The 9-MeA coordinates through the N1 and N3 positions to two Ag(I) centres. The backbone of the polymer is a helical [Ag1-N3a-C2a-N1a]∞ chain. This complex was the first example reported of simultaneous binding of three metal ions to a neutral adenine, via N7 (Pt(II)), N1 (Ag(I)) and N3 (Ag(I)).23 Coordination polymers containing Cu(II) have been reported for adenine and guanine, respectively.24 These compounds, prepared by reacting Cu(II) ions with alkyldiamine-tethered adenine or guanine, were isolated as helical polymers. In both

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Figure 4.22 Representation of the helical structure of the [{(NH3)2Pt(9-MeA-N7)(9-MeHxN7)}Ag(NO3)(H2O)](ClO4)(NO3) coordination polymer

polymers the metal centres adopt square pyramidal geometry, however the details of the metal-nucleobase binding are quite different. In the guanine-containing material, {[Cu(N7-Et-G-en)(H2O)2]2+}n, N7-binding is observed, whereas in the adenine derivative, {[CuCl(N7,N3-Et-A-en]+}n, both N7 and N3 atoms are bound to the metal centre. This difference in coordination has a dramatic effect on the shape of the polymer chains with essentially parallel guaninyl residues in the former case compared to an interplanar angle of ∼74 ° for adjacent bases in the latter. The helical arrangement of the former compound with a pitch of ∼12 Å and Cu···Cu (Figure 4.23) distances around the periphery of the helical structure of 7.0 Å is similar to metrical parameters in native duplex DNA and raises the interesting possibility of metal–nucleobase polymers acting as antisense agents. Srivantsan and coworkers have described the catalytic activity of nucleobase– copper polymeric frameworks.25 Studies pertaining to prebiotic catalysis and the chemical origin of life have attracted considerable attention over the years.26 In this context, the possible assistance of mineral surfaces for biological polymerization reactions has been pointed out.27 It has been noted that Cu(II) ions assist the binding of nucleotides to mineral surfaces. The structure of 9-(4-vinylbenzyl)adenine–copper polymer, which shows the copper ions coordinated to N1 and N7 positions of adenine (Figure 4.24), mimics the periodicity of single-stranded nucleic acids.25 The cross-linking of the polymeric chains of this complex with ethyleneglycol dimethacrylate or 1,4-divinylbenzene ensured its insolubility in aqueous solutions and, therefore, allowed biological studies to be performed, which revealed that these

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Figure 4.23 Schematic illustration of the helical chain observed in the Cu-guanine polymer [Cu(H2O)2(N7-G-Et-en)]2+

Figure 4.24 Zig-zag structure of the 1D coordination polymer of an adenine-derivative copper complex

Figure 4.25 1D Cu(II) adenine glycine polymer

compounds catalyse nucleic acid cleavage in heterogeneous conditions. This fact suggests that such polymeric structures could serve as models of the cooperative interaction between nucleic acid constituents and metal ions for prebiotic catalysis.25 A monodimensional copper(II) infinite zigzag chain polymer, with coordinated adenine and glycine moieties is represented in Figure 4.25. In this polymer the

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Figure 4.26 Scheme of the structure of the 1D coordination polymer [Cd(6-MP−)2]n

adenine molecule acts as bridging ligand in a zigzag manner. The copper ions coordinate via N1 and N9 to propagate the polymeric chain along the crystallographic b-axis. Again the structure resembles the expected periodicity of single-stranded nucleic acids. A water suspension of this insoluble complex exhibits not only facile chemical nuclease activity for DNA by an oxidative mechanism, but can also be reused, keeping its scission activity unchanged.28 The direct reaction between Cd(NO3)2, and 6-MP (6-mercaptopurine) (pH 4.6) leads to formation of the coordination polymer [Cd(6-MP−)2·2H2O]n (Figure 4.26).29 Its X-ray structure consists of infinite chains of cadmium atoms octahedrally coordinated by four S6 and two N7 atoms of the m-S6,N7:S6-mercaptopurinate ligands. The metal-to-metal distance within the chain is 3.92 Å. Addition of 0.1 M NaOH and a 0.1 M water solution of Ca(NO3)2 to a suspension of this polymer leads to further deprotonation of the 6-MP− ligand and the subsequent formation of the coordination polymer [Cd(6-MP2−)2]n[Ca(H2O)6]n.30 Its structure consists of parallel 2 n− 2 n+ anionic chains of [Cd (6-MP 2 − )2 ]n and cationic chains of [Ca ( H 2O )6 ]n which are joined together by means of electrostatic interactions and hydrogen bonds. The 1D anionic polymer is based on stacked centrosymmetric [Cd(6-MP2−)2] entities, which are linked to each other through Cd–S bonds, forming a 1D chain. The ligand coordinates to the cadmium as in the former case. The distance between metal centres of consecutive entities is 4.14(1) Å. An interesting issue is the different shape of the two polymeric chains and the prevalence of the trans disposition in the [Cd(6-MP−)2] entities. The structural analogy between [Cd(6-MP−)2] and that proposed for M-DNA (see Section 4.5), motivated studies of the electrical properties of single molecular strands. In spite of [Cd(6-MP−)2] behaving as an insulator, the work opens a new strategy towards the possible use of DNA-derived coordination polymers as molecular wires.31 The reaction of tetrakis(m-trifluoroacetamidato)dirhodium(II) with cytosine in methanolic/aqueous solution yields a zigzag 1D structure in which the cytosine molecule bridges the axial positions of the different centrosymmetric [Rh2(CF3CONH)4] units through N3 and O2 donor atoms (Figure 4.27).32 The Rh– N3 axial bonding is accompanied by the formation of two intramolecular hydrogen bonds, one between N4 of the base and the amidato oxygen and the other between O2 of the base and the amidato NH. The Rh–O2 coordinative bond is also reinforced by a hydrogen bond established between the cytosine N1 atom and the amidato oxygen atom of a trifluoroacetamidato bridge.

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Figure 4.27 Zigzag 1D structure of [Rh2(CF3CONH)4(1-MeC-O2,N3)]n

Figure 4.28 Schematic illustration of [Cd4(N7-G-Et-en)4(H2O)4(SO4)2] highlighting the 1D porous structure generated through sulfate bridging of G-quartets

Nucleobases and/or Anions as Bridging Ligands Houlton and coworkers were able to isolate crystals of a polymeric compound based on tetrameric [Cd4(G-Et-en)4(H2O)4] entities, which are assembled to form waterlined pores by bridging sulfate anions11. Within the tetrameric assembly each Cd(II) atom is coordinated by a tethered diamine function and N7 of an adjacent guanine moiety. This arrangement leads to a cylindrical (square) polymer containing an ′ (Figure 4.28) as we have described internal channel with a diameter of about 12 Å before see Section 4.2.2).

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Figure 4.29 Linear chain of compound [Cd2(N6-benzyladenine)2(µ-Cl)4Cl2]n

Figure 4.30 Polymeric chain of compound {[HgCl2(1,9-dimethyl-guanine-N7)]·2HgCl2}n

Stanley and coworkers have reported the structures of two monodimensional polymers of N6-substituted adenine derivatives (N6-furfuryladenine and N6benzyladenine) coordinated to Cd(II)33. Both ligands present interesting properties as plant growth stimulants.34 In these structures, the Cd(II) ions adopt a distorted octahedral geometry, with coordination to N9 and five chloride ions. Successive Cd(II) ions are bridged by two chloride ions (Figure 4.29). Lippert and coworkers have synthesized a 1D polymeric Hg(II)-nucleobase complex from the guanine derivative (1,9-DiMeG = 1,9-dimethylguanine), by the reaction of HgCl2 in a water/alcohol mixture (Figure 4.30). The Hg(II) ion is coordinated to one guanine (via the N7 position) and three Cl ligands, two of which act as bridging ligands. The polymeric structure is cocrystallized with HgCl2 molecules.35 Two isomorphous Cd polymers of chelate-tethered adenine derivatives have been characterized. Reactions of Cd(II) salts with A-Et-enH·Cl in water yielded material with the stoichiometry [{Cd2X6(A-Et-en)2}n] (X = Cl and Br). Each compound contains two unique Cd(II) ions: a central cadmium which sits on a centre of symmetry and is coordinated in an octahedral manner by four bridging halide anions and the N7 of two trans-arranged adenines. The second cadmium adopts a distorted octahedral geometry with three halide ligands, two of these bridging and

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Figure 4.31 Schematic illustration of the polymeric chains in [µ-{CdBr(N3,N7-A-Eten)}2CdBr4], (where X = Cl or Br) generated by adeninyl bridging of trimetallic units

the third terminal. The remaining coordination sites are occupied by the nitrogen atoms of the diamine group and N3 of adenine (Figure 4.31).36 García-Terán and coworkers have found a strategy for the design of 1D oxalatocomplexes which have been extended to the synthesis of infinite MII-oxalato chains with pendant purine, [M(m-ox)(H2O)(pur)]n (M = Cu, Co and Zn), and adenine, {[(M(m-ox)(H2O)(ade)]·2(ade)(H2O)]}n (M = Co and Zn), nucleobases.37 The polymeric chains consist of cis-[M(H2O)(L)]2+ units bridged by bis-bidentate oxalato ligands with M···M intrachain distances in the range 5.23–5.57 Å (Figure 4.32a, b). The metal atoms are six-coordinate with four oxalato oxygen atoms, one water molecule and one nitrogen atom from a terminal nucleobase, building distorted octahedral MO4OwN surroundings. The purine ligand is bound to the metal atom through the most basic imidazole N9 atom, whereas for the adenine nucleobase the minor groove site N3 is the donor atom. The nucleobases are projected outside the zigzag chain in both structures, setting up a complex network of noncovalent interactions. The interplanar distance of 7.52 A between two consecutive parallel adenine ligands on the same side of the chain permits the insertion of free adenine molecules between, and approximately parallel to, the coordinated adenines, with a mean interplanar distance of 3.22 A. This value is somewhat shorter than that found between adjacent bases in the B-DNA double helix (3.4 A). Additionally, a second free adenine interacts with the coordinated adenine molecule through the donor and acceptor sites of both Watson–Crick (N1 and N6 atoms) and Hoogsten (N7 and N6 atoms) edges to give a zigzag array with the intercalated free adenine forming a strong N9-H···N3 hydrogen bond. Furthermore the cohesiveness of the supramolecular architecture is ensured by three additional hydrogen bonds (N9-H···Ow, OwH···N3 and C8-H···Ow) (Figure 4.32c). These compounds are rare examples of the coexistence of high dimensional M–nucleobase covalent and supramolecular networks.

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Figure 4.32 Fragments of the 1D polymeric complex: (a) [M(µ-ox)(H2O)(pur)]n and (b) {[(M(µ-ox)(H2O)(ade)]·2(ade)(H2O)]}n; (c) hydrogen-bond network established among the coordinated and free adenine molecules

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4.3.2 2D Coordination Polymers Nucleobases and Anions as Bridging Ligands. The examples of 2D covalent frameworks, based on the coordinative properties of the nucleobases, are rare. No examples with only bridging nucleobase bidimensional networks have been described to date. The two cases encountered in the CSD database exhibit cooperative bridging of a nucleobase and of an auxiliary anion. Lippert et al. reported the first of these two cases in which trans-[Pt(CH3NH2)2(cytosinatoN1)(cytosinato-N3)] molecules are cross-linked by CuII ions through the still available N3 and N1 positions of the cytosine ligands to give a 1D coordination polymer. These ···Pt-cytosinato-Cu-cytosinato-Pt··· chains are further cross-linked by means of double m-hydroxo bridges between CuII atoms belonging to adjacent chains leading to the overall corrugated 2D crystal structure (Figure 4.33).38 The 2D extended solid-state structure can also be described as fused rings comprised of 4 Pt and 6 Cu metal centres. Verma and Purohit recently reported a 2D framework constructed from 9allyladenine and silver nitrate.39 Its asymmetric unit contains distorted tetrahedral

Figure 4.33 Section of the 2D heteronuclear [trans-[Pt(CH3NH2)2(µ-cytosinato-N1,N3)(µcytosinato-N3,N1)Cu(OH)]ClO4·1.2H2O coordination complex

Infinite Architectures 119

Figure 4.34 2D framework of [Ag(µ-9-allyladenine-N1,N7)(µ-NO3)(NH3)]n compound

and square pyramidal silver atoms bridged by adenine molecules. The adenine coordinates through N1 to the tetracoordinated metal centre and through N7 to the pentacoordinated one. This generates a helical strand in which the two metal centres alternate. The helical coordination chain possesses two adenine residues per turn ′ . The crystal structure is completed by the presence of bridging with a pitch of ∼2.5 Å nitrate counterions binding to the helical strands via silver ions. The authors were able to deposit this compound on highly oriented pyrolitic graphite retaining the solid-state structure, as confirmed by noncontact mode AFM micrographs. Deposition on alternative substrates such as Si(100), or mica, lacked obvious pattern formation, suggesting that known adenine–graphite interactions40 play a role in the preferential orientation observed on graphite. The authors describe these results in the context of efforts to functionalize surfaces for advanced nanoscale applications by means of shape-persistent coordination motifs (Figure 4.34).41–42 4.3.3 3D Coordination Polymers Nucleobases as Bridging Ligands. In spite of the great number of potential donor atoms of the nucleobases, especially in the case of adenine, examples of 3D coordination polymers containing bridging nucleobases are as of today limited to just two different types of structures: one 3D porous coordination polymer and a family of isostructural luminiscent 3D polymers. The first example is the {[Cu2(m-ade)4(H2O)2][Cu(ox)(H2O)]2·14H2O}n compound which shows a 3D coordination network, where the Cu(II) centres are bridged by tridentate m-N3,N7,N9-adeninato ligands (Figure 4.35).43 The crystal structure is constructed of two distinct types of neutral building blocks: windmill shaped dimeric [Cu2(m-ade)4(H2O)2] units and mononuclear [Cu(ox)(H2O)] entities. The deprotonation of the adenine molecules allows the N9 position to act as a donor and in this

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Figure 4.35 Repetitive unit of the {[Cu2(µ-ade)4(H2O)2][Cu(ox)(H2O)]2·14H2O}n

3D

porous

coordination

polymer

way provides additional binding, linking the [Cu(ox)(H2O)] entities. This is also observed in the discrete hexameric {(H2O)2Cu2(m3-(A)4[Cu(oda)(H2O)]4}·6H2O (oda = oxydiacetato) entity (see Section 4.2.3), in which the central dimeric core is bonded to four monomeric units.16 In the structure, the [Cu(ox)(H2O)] entities presents two available coordination positions, which allow each of these moieties to be linked to two dimeric [Cu2(made)4(H2O)2] cores, leading to a 3D coordinate framework. This self-assembly process directed by the metal–adeninate linkages, creates nanoscale channels with a diameter of about 13 Å (40.4% of the crystal volume). Many guest water molecules are trapped within the parallel nanotubes. The structure remains unaltered when the water molecules are removed. Verma and coworkers have reported the structures of an argentite adenyl homopolymer, with interesting luminescent properties. The 3D polymeric compound formed by 9-allyladenine (9-AA), contains a metallomacrocyclic quartet, where N1, N3 and N7 nitrogens are coordinated to silver ions (Figure 4.36). Four silver ions are located at each corner of a distorted rectangle (interatomic distances for the two sides of the rectangle: 7.71 and 6.95 Å). The structure is composed of four noninterpenetrating parallel strands, two 1D left-handed and two 1D righthanded helices. This unusual coordination mode is evocative of DNA quadruplex formation with four purine residues. This type of structure may lead to the generation of metallomacrocyclics with different pore sizes suitable for gas storage and

Overview of Metal Coordination Sites

121

Figure 4.36 Metallomacrocyclic quartets linking chiral chains of [Ag(µ-allyladenine-N1,N3)] to afford the resulting 3D coordination polymer

novel photoactive materials.44 More recently, Verma has published five new compounds of the same family based on silver, modified adenine moieties and counterions: 9-allyladenine-silver perchlorate, 9-allyladenine-silver tetrafluoroborate, 9-allyl-6-chloropurine-silver nitrate, 9-propyladenine-silver nitrate and 9-propyl-6chloropurine-silver nitrate.45

4.4 Overview of Metal Coordination Sites and Base–Base Hydrogen Bonding In this section, instead of a detailed discussion on selected examples, we provide a brief summary of the supramolecules that have been formed exclusively through coordinate bond formation involving the nucleobases and, secondly, supramolecules which self-assembly through H-bonding among the nucleobases acting as ligands to metal centres. Therefore, this section will provide a general overview of the current situation in this field without going into specific details of structures and properties. More than 600 metal–nucleobase complexes have been reported to date. This is sufficient for a statistically significant study of the diverse metal binding patterns, assemblages of hydrogen bonding, and, in particular, self-pairing of the existing metal–nucleobase complexes. This study was performed via the Cambridge Crystallographic Data Base.46

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Figure 4.37 Schematic representation of H-bonding patterns involving cytosine.

Figure 4.38 Schematic representation of H-bonding patterns involving thymine/uracil.

Self-pairing of nucleobases requires hydrogen-bond interactions between edges of different nucleobases: (i) minor groove edge, Hoogsteen and Watson–Crick edges in purines; (ii) sugar and Watson–Crick edges in pyrimidines. Evidently, coordination of a metal to a nucleobase hinders the pairing of that site (edge), although the pairing of other edges can be favoured, for example through pKa alterations.47–53 A graphic scheme of the existing self-pairing of metal-nucleobase complexes patterns is depicted in Figures 4.37–4.40. The dataset may be categorized from the viewpoint of the number, and location, of coordinated metal ions. It is apparent that there is a wide variety of metal ion binding modes to the different nucleobase sites (Table 4.1). For statistical reasons 5-MeC was considered as C, and N-alkylated bases as their unalkylated ones. The format in Table 4.1 is as follows: the location (G, N1), for instance, corresponds with 21(13), where 21 is the number of examples containing a metal coordinated at the N1 position of guanine and 13 is the number of molecules where N1 is the only metal-coordinated position. The type of transition metal was not considered, but is usually Pt, Pd, Ag, Cu or Hg. As evident from Table 4.1, examples of metal ions bonded to all accessible coordination sites of the different nucleobases have been found. Even endocyclic carbon atoms, such as the C5 of cytosine17 and uracil, and C8 of adenine54 and guanine,55 can substitute their protons by metal ions. Adenine can be considered the most versatile nucleobase, with many examples of monometallated derivatives featuring binding at different sites. Guanine is an extreme case, with 184 monometallated complexes found with N7-coordination and

Overview of Metal Coordination Sites

123

Figure 4.39 Schematic representation of H-bonding patterns involving guanine

Figure 4.40 Schematic representation of H-bonding patterns involving adenine Table 4.1 Examples containing all the possible metal coordination sites N1

N2/O2

C

11 (7)

19 (5)

N3

N4/O4

U

17 (6)

14 (0)

77 (25)

47 (2) 30 (0)

199 (131) 70 (11)

C5

N6/O6

N7

C8

N9

1 (0) 12 (12)

T

8 (7)

2 (0)

68 (37)

G

21 (13)

1 (0)

4 (0)

8 (0)

A

55 (13)

41 (20)

44 (7)

198 (184) 2 (2) 15 (11) 129 (65)

1 (1) 56 (26)

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only 26 complexes with coordination found at other sites. For uracil, however, even though the majority of the reported cases are N3-coordinated, the number of complexes with metal coordinated to other positions is considerable. One of the essential points in the interpretation of Table 4.1 is related to mechanistic aspects. Direct coordination of a metal to a nucleobase usually involves the preferred sites: N7 in purines, N3 in pyrimidines. After coordination, the metal can migrate to other positions, or the nucleobases can even undergo a pKa alteration, making it easier for further coordination by another metal ion. Thus, multiple coordination favours the coordination of a metal ion to an initially nonpreferred site. In Table 4.1 all the cases of multimetal complexes were counted as individual entities for each site. Though it is possible to form chelate binding modes for nucleobases, and these have been proposed in earlier literature, there are remarkably few well-characterized examples. The most abundant cases are for the adenine nucleobase, which shows two different chelating modes: N1,N6 and N6,N7. Two examples, both with molybdenum,56 are known for the N1,N6 mode, and several examples with different metals for the latter: platinum (3), copper (2), ruthenium (5), rhodium (3), iridium (2).57 Metal chelation has also been described for the other nucleobases, with the exception of thymine. For guanine, only one example has been found, involving zinc bound through N1,O6.58 This is perhaps an unexpected feature given the large number of guanine complexes characterized. The number of examples containing pyrimidine bases is also very small. One example is known for uracil, in which a copper ion is bound to the base at both N1 and O2.59 Two chelating modes were found for cytosine: O2,N3 and N3,N4, comprising six different cases involving cadmium (4), copper (1), and rhodium (1).60 The direct utilization of DNA base pairing to control the assembly of metal complexes has been realized by Houlton. This was achieved by extending ligand functionalization to include two complementary nucleobases (Figure 4.41).61 Sequential alkylation of 1,2-dithioethane using 2-chloroethyl-9-adenine and 3-chloropropyl1-thymine yields the AT-containing dithioether. Reaction with either [PdCl2(MeCN)2] or K2[PtCl4] gives the isostructural complex cations, [MCl(N3-A-SS-T)]+ (M = Pd(II) or Pt(II)), in which the dithioether strand is coordinated through the S-atoms of the thioether and N3 of adenine. In the solid state the complex cations form infinite chains of Hoogsteen base pairs (T-N3···A-N7 separation 2.89 Å), which is the most thermodynamically stable pattern for these bases. It was also found that the effect of metal ion binding to the adeninyl group is to reduce the hydrogen bonding between the A-T pair in solution compared to the free ligand.62 This approach provides a range of possibilities for the development of supramolecular syntheses, which take advantage of both coordinate bonding and complementary hydrogen bonding.

4.5 M-DNA The use of DNA as an electrically conductive material, with self-organization properties able to produce molecular nano-architectures, has motivated extensive

M-DNA

125

Figure 4.41 (a) Stepwise synthesis of a self-complementary adeninyl–thymine containing thioether, A-SS-T. (b) Scheme illustrating the structure of the adeninyl–thymine-containing complexes [MCl(N3-A-SS-T)]+ (M = Pd(II) or Pt(II)). (c) Scheme illustrating the Hoogsteen base pairing observed in the crystal structure of [MCl(N3-A-SS-T)]+

research in recent years.63 However, the direct electrical characterization of DNA has been controversial64 since this molecule has been described variously as a conductor65 and an insulator.66 Currently it is widely accepted that it behaves as an insulator from 30 nm lengths up.67 Several approaches have been attempted to try to modify the electronic structure of DNA in order to increase conductivity.63 A new molecular wire candidate based on DNA is so-called M-DNA.68 This type of DNA consists of a complex formed between divalent metal ions (Zn2+, Ni2+ and Co2+) and duplex DNA (Figures 4.42 and 4.43). M-DNA is formed at high pH (7.5–9.0) and the model proposed consists of a structure in which the metal cations replace the imino protons involved in H-bonds between A-T and G-C base-pairs (Figure 4.43). As a result, the metal ions are located in the interior of the helix resulting in metal–metal distances close to 3.4 Å, corresponding to the value of the typical pstacking between nucleobases in B-DNA. However, despite preliminary NMR data consistent with liberation of the protons inside the strand, the exact position of the metals remains unclear. Interestingly, M-DNA converts back to B-DNA either when the pH of the solution decreases or when chelating agents such as EDTA are added. Among several interesting physical features, Lee et al. have reported that DNA strands modified with fluorescent probes, fluorescein and rhodamine, show a strong quenching of the fluorescence upon M-DNA formation. They suggested that this effect could be caused by a fast electron transfer.69 However, the electrical charac-

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Supramolecular Chemistry of Metal–Nucleobase Complexes

Figure 4.42 Preferred base-pairing schemes for M-DNA in which imino protons of T and G are replaced by metal cations

Figure 4.43 Scheme of the proposed structure of M-DNA

terization of M-DNA is again controversial. Thus, while Rakitin et al. reported metallic-like conduction in a 15 mm long bundle of M-DNA,70 recent new direct atomic force microscope measurements show that in spite of the DNA length being greatly reduced when M-DNA is formed, the conductivity is not enhanced.71 Probably, these results are related to the low stability of the M-DNA structure, as mentioned above. In order to understand the experimental results of the electronic properties of M-DNA, several theoretical calculations have been performed. Di Felice et al. found that Zn(II) does not efficiently hybridize with the HOMO orbital of the native G-C pair, and suggested that the model proposed by Lee should be considered with caution.72 DFT calculations carried out by Alexandre et al. on a new M-DNA structure with the metal ions located in the major groove of DNA replacing the imino protons, but in a pseudo-tetrahedral environment instead of the square planar suggested by Lee,73 indicate that the conductivity of Zn-DNA, Co-DNA and Fe-DNA could increase significantly. The improved stacking effect observed in Zn-DNA suggested a new possible route to increase DNA conductivity. Finally,

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studies performed by Fuentes-Cabrera et al. pointed out that neither planar nor nonplanar models have electronic properties that correlate with M-DNA (M = Zn) or could give an explanation for a measured conductivity for M-DNA in terms of metal cation d-states.74 The enhanced conductivity could be due to a significant decrease of the HOMO–LUMO gap of some forms of M-DNA as compared to that of DNA, but more research is needed to understand whether further reduction in the HOMO–LUMO gap takes place upon metal ion binding to base pairs. Despite the above-mentioned difficulties in the characterization of M-DNA, potential use as molecular wires in nanocircuits makes M-DNA, or synthetic analogues, promising materials for nanotechnological applications.

4.6 Conclusion The aim of this chapter has been to provide an overview of the supramolecular aspects of metal–nucleobase chemistry. Much of the small molecule coordination chemistry of DNA bases is well established through the efforts of numerous groups. However, the organization of molecules into functional systems, with specific tailor-made properties, remains a challenge. The hugely anticipated development of molecular-based devices, along with the shift from top-down to bottom-up assembly, still requires key contributions from a range of disciplines. It is perhaps by combining the unique properties of metal ions, e.g. switchable oxidation states, electronic and magnetic effects etc., with the diverse ligand-binding modes of nucleobases, that those important breakthroughs may be made. The bifunctionality of the building blocks of life, multiple metal–ion binding allied with complementary hydrogen bonding, offers a powerful means by which to organize molecular systems. From the breadth of complex types discussed herein, it is obvious that metal ion–nucleobase interactions will continue to warrant research for many years to come.

Acknowledgements Financial support by the Ministerio Español de Educación y Ciencia, Comunidad de Madrid (projects no. MAT2007-66476-C02-02, MAT2005-03047, NAN200409183-C10-06, CTQ2006-027185-E, and S-0505/MAT/0303) and European Community (FP6-029192) is gratefully acknowledged.

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38. 39. 40.

41. 42. 43. 44. 45. 46. 47. 48. 49.

Supramolecular Chemistry of Metal–Nucleobase Complexes Synthesis and scanning probe microscopy characterization; Inorg. Chem., 2006, 45, 7642–7650. Olea, D.; Alexandre, S.S.; Amo-Ochoa, P.; Guijarro, A.; de Jesus, F.; Soler, J.M.; de Pablo, P.J.; Gómez-Herrero J.; Zamora, F.; From coordination polymer macrocrystals to nanometric individual chains; Adv. Mater., 2005, 17, 1761–1764. Aoki, K; Salam, M.A.; Interligand interactions affecting specific metal bonding to nucleic acid bases: the tetrakis(µ-trifluoroacetamidato)dirhodium(II)-cytosine system. Crystal structures of [Rh-2(CF3CONH)4(cytosine)] and [Rh-2(CF3CONH)4(1methylcytosine)2]·2H2O; Inorg. Chim. Acta, 2001, 316, 50–58. Stanley, N.; Muthiah, P.T., Lugar, P.; Weber, M.; Geib, S.J.; Metal-nucleobase interactions: Interplay of coordination and hydrogen bonding in cadmium(II) complexes of N6substituted adenines; Inorg. Chem. Comm., 2005, 8, 1056–1059. Pattabhi, V.; Plant-growth regulators – their structure and interactions; Curr. Sci. India, 1990, 59, 1228–1235. Menzer, S.; Hillgeris, E.C.; Lippert, B.; Preparation, HgCl2 coordination to guanine derivatives. Structural and spectroscopic studies on the interactions with 9-ethylguanine, 6,9dimethylguanine and 2-amino-6-methoxy-9-methylpurine; Inorg. Chim. Acta, 1993, 211, 211–226. Amantia, D.; Shipman, M.A.; Price, C.; Elsegood, M.R.J.; Clegg, W.; Houlton, A.; Conserved supramolecular architecture in polymetallic nucleobase complexes; Inorg. Chim. Acta, 2006, 359, 3515–3520. García-Terán, P.; Castillo, O.; Luque, A.; García-Couceiro, U.; Beobide, G.; Román, P.; Supramolecular architectures assembled by the interaction of purine nucleobases with metal-oxalato frameworks. Non-covalent stabilization of the 7H-adenine tautomer in the solid-state; Dalton. Trans., 2006, 902–911. W. Brüning, W.; Freisinger, E.; Sabat, M.; Sigel, R.K.O.; Lippert, B.; N1 and N3 linkage isomers of neutral and deprotonated cytosine with trans-[(CH3NH2)2Pt(II)]; Chem. Eur. J., 2002, 8, 4681–4692. Purohit, C.S.; Verma, S.; Patterned deposition of a mixed-coordination adenine-silver helicate, containing a pi-stacked metallocycle, on a graphite surface; J. Am. Chem. Soc., 2007, 129, 3488–3489. (a) Ortman, F.; Schmidt, W.G.; Bechstedt, F.; Attracted by long-range electron correlation: Adenine on graphite; Phys. Rev. Lett., 2005, 95, 186101–186104; (b) Sowerby, S.J.; Cohn, C.A.; Heckl, W.H.; Holm, N.G.; Differential adsorption of nucleic acid bases: Relevance to the origin of life. Proc. Nat. Acad. Sci., 2001, 98, 820–822. Barth, J.V.; Costantini, G.; Kern, K.; Engineering atomic and molecular nanostructures at surfaces; Nature, 2005, 437, 671–679. Theobald, J.A.; Oxtoby, N.S.; Phillips, M.A.; Champness, N.R.; Beton, P.H.; Controlling molecular deposition and layer structure with supramolecular surface assemblies; Nature, 2003, 424, 1029–1031. García-Terán, J.P.; Castillo, O.; Luque, A.; García-Couceiro, U.; Román, P.; Lezema, L.; An unusual 3D-coordination polymer based on bridging interactions of the nucleobase adenine; Inorg. Chem., 2004, 43, 4549–4551. Purohit, C.S.; Verma, S.; A luminescent silver-adenine metallomacrocyclic quartet; J. Am. Chem. Soc., 2006, 128, 400–401. Purohit, C.S.; Mishra, A.K.; Verma, S.; Four-stranded coordination helices containing silver-adenine (purine) metalloquartets; Inorg. Chem., 2007, 46, 8493–8495. Cambridge Crystallographic Database, November 2006. Sigel, H., Complex Formation of Nucleic Bases with Cu2+. Acidity of N-1-protons in inosine guanosine uridine and thymidine 5′-triphosphate and in their Cu2+ complexes; Eur. J. Biochem., 1968, 3, 530–537. Sigel, H.; Calase and peroxidase activity of Cu2+ complexes; Angew. Chem., 1969, 81, 161–171. Kastner, M.E.; Coffey K.E.; Clarke M.J.; Edmonds S.E.; Eriks K.; Structural correlation and metal-ion movement in stable pentaammineruthenium(III)-hypoxanthine complexes; J. Am. Chem. Soc., 1981, 103, 5749–5752.

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50. Clarke, M.J.; Taube, H.; Pentaammineruthenium-guanine complexes; J. Am. Chem. Soc., 1974, 96, 5413–5419. 51. Chu, G.Y.H.; Mansy, S.; Duncan, R.E.; Tobias, R.S.; Heavy-metal nucleotide interactions. 12. competitive reactions in systems of 4 nucleotides with cis-diammineplatinum(II) or trans-diammineplatinum(II) – Raman difference spectrophotometric determination of relative nucleophilicity of guanosine, cytidien, adenosine, and uridine monophosphates as well as analogous bases in DNAM; J. Am. Chem. Soc., 1978, 100, 593–606. 52. Sigel, H.; Lippert, B.; Sigel, H; Lippert, B.; The effects of N7-coordinated cis-diammineplatinum(II) on the acid-base properties of guanine derivatives; Pure Appl. Chem., 1998, 70, 845–854. 53. Roitzsch, M.; Añorbe, M.G.; Sanz Miguel, P.J.; Müller, B.; Lippert, B., The role of intramolecular hydrogen bonding on nucleobase acidification following metal coordination: possible implications of an ‘indirect’ role of metals in acid-base catalysis of nucleic acids; J. Biol. Inorg. Chem., 2005, 10, 800–812. 54. Price, C.; Elsegood, M.R.J.; Clegg, W.; Rees, N.H.; Houlton, A.; Evidence for directed metalation: A structural intermediate in the formation of a novel C-bound adenine complex of ruthenium; Angew. Chem. Int. Ed., 1997, 36, 1762–1764. 55. Neidle, S.; Kuroda, R.; Broyde, S.; Hingerty, B.E.; Levine, R.A.; Miller, D.W., Evans, F.E.; Conformation of the C8 substituted guanine adduct of the carcinogen acetylaminofluorene: model for a possible Z-DNA modified structure; Nucl. Ac. Res., 1984, 12, 8219. 56. Kuo, L.Y.; Kanatzidis, M.G.; Marks, T.J.; Metallocene antitumor agents – unusual Mo(eta5-C5H5)2Cl2 nucleotide nucleobase aqueous coordination chemistry; J. Am. Chem. Soc., 1987, 109, 7207–7209. 57. For example: (Pt) ref 4; (Cu) ref 16b; (Ru) Korn, S.; Sheldrick, W.S., pH-dependent competition between k(2)N-7,O(P) macrochelation and µ-N-1,N-7 oligomer formation for (eta(6)-arene)Ru-II complexes of adenosine and guanosine 5′-mono-, -di- and -triphosphates; J. Chem. Soc., Dalton Trans., 1997, 2191–2199.; (Rh) ref 8; (Ir) Annen, P.; Schildberg, S.; Sheldrick, W.S., (eta(5)-Pentamethylcyclopentadienyl)iridium(III) complexes of purine nucleobases and nucleotides: a comparison with (eta(6)-arene) ruthenium(II) and (eta(5)-pentamethylcyclopentadienyl)rhodium(III) species; Inorg. Chim. Acta, 2000, 307, 115–124. 58. Badura, D.; Vahrenkamp, H.; Pyrazolylboratozinc complexes of nucleosides and nucleoside analogues; Eur. J. Inorg. Chem., 2003, 3723-3730 59. Fridman, Y.D.; Tsoi, E.P.; Levina, M.G.; Synthesis and properties of copper orotates and copper mixed-ligand compounds with orotic-acid and amino-acids; Koord. Khim. (Russ.) (Coord.Chem.), 1984, 10, 469–474. 60. For example: (Cd) Shiba, J.K.; Bau, R.; Crystal and molecular-structure of the monoclinic form of [Cd(5′-CMP)(H2O)]N1; Inorg. Chem., 1978, 17, 3484–3488; (Cu) Szalda, D.J.; Kistenmacher, T.J., Transition-metal-nucleoside complexes – crystal and molecularstructure of (glycylglycinato)(cytidine)copper(II) dihydrate; Acta Cryst., 1977, B33, 865– 869; (Rh) Smith, D.P.; Olmstead, M.M.; Noll, B.C.; Maestre, M.F.; Fish, R.H.; Bioorganometallic chemistry 2. Synthesis and structural studies of the reactions of a nucleobase, 1-methylcytosine, with a (eta-5-pentamethylcyclopentadienyl)rhodium aqua complex; Organometallics, 1993, 12, 593; the second modus N3,N4 was found in four cases of platinum (3): Beyerle-Pfnur, R.; Schollhorn, R.; Thewalt, U.; Lippert, B.; J. Chem. Soc. Chem. Commun., 1985, 1510; Schollhorn, R.; Beyerle-Pfnur, R.; Thewalt, U.; Lippert, B., Unusual 4-membered chelate rings of Pt-IV with a cytosine nucleobase; J. Am. Chem. Soc., 1986, 108, 3680–3688; and molybdenum (1) Kuo, L.Y.; Kanatzidis, M.G.; Sabat, M.; Tipton, A.L.; Marks, T.J., Metallocene antitumor agents – solution and solid-state molybdenocene coordination chemistry of DNA constituents; J. Am. Chem. Soc., 1991, 113, 9027–9045. 61. Price, C.; Mayeux, A.; Horrocks, B.R.; Clegg, W.; Houlton, A.; Self-complementary metal complexes containing a DNA base pair; Angew. Chem. Int. Ed., 2002, 41, 1047. 62. Amantia, D.; Price, C.; Shipman, M.A.; Elsegood, M.R.J.; Clegg, W.; Houlton, A.; Minor groove site coordination of adenine by platinum group metal ions: Effects on basicity, base pairing, and electronic structure; Inorg. Chem., 2003, 42, 3047–3056.

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63. Carell, T.; Behrens, C.; Gierlich, J.; Electrontransfer through DNA and metal-containing DNA; Org. Biomol. Chem., 2003, 1, 2221–2228. 64. Porath, D.; Cuniberti, G.; Di Felice, R.; Charge transport in DNA-based devices; Topics Curr. Chem., 2004, 237, 183–227. 65. Kasumov, A.Y.; Kociak, M.; Guéron, S.; Reulet, B.; Volkov, V.T.; Klinov, D.V.; Bouchiat, H. Proximity-induced superconductivity in DNA; Science, 2001, 291, 280–282. 66. de Pablo, P.J.; Moreno-Herrero, F.; Colchero, J.; Gomez-Herrero, J.; Herrero, P.; Baro, A.M.; Ordejon, P.; Soler, J.M.; Artacho, E.; Absence of dc-conductivity in lambda-DNA; Phys. Rev. Lett., 2000, 85, 4992–4995. 67. Porath, D.; Cuniberti, G.; Di Felice, R. Charge transport in DNA-based devices. Topics Curr. Chem., 2004, 237, 183–227. 68. Aich, P.; Labiuk, S.L.; Tari, L.W.; Delbaere, L.J.T.; Roesler, W.J.; Falk, K.J.; Steer, R.P.; Lee, J.S.; M-DNA: A complex between divalent metal ions and DNA which behaves as a molecular wire; J. Mol. Biol., 1999, 294, 477–485. 69. Wettig, S.D.; Wood, D.O.; Aich, P.; Lee, J.S.; M-DNA: A novel metal ion complex of DNA studied by fluorescence techniques; J. Inorg. Biochem. 2005, 99, 2093–2101. 70. Rakitin, A.; Aich, P.; Papadopoulos, C.; Kobzar, Y.; Vedeneev, A.S.; Lee, J.S.; Xu, J.M.; Metallic conduction through engineered DNA: DNA nanoelectronic building blocks; Phys. Rev. Lett., 2001, 86, 3670–3673. 71. Moreno-Herrero, F.; Herrero, P.; Moreno, F.; Colchero, J.; Gomez-Navarro, C.; GomezHerrero, J.; Baro A.M.; Topographic characterization and electrostatic response of MDNA studied by atomic force microscopy; Nanotechnol., 2003, 14, 128–133. 72. Di Felice, R.; Calzolari, A.; Zhang, H.; Towards metalated DNA-based structures; Nanotechnol., 2004, 15, 1256–1263. 73. Alexandre, S.S.; Soler, J.M.; Seijo, L.; Zamora, F.; Geometry and electronic structure of M-DNA (M = Zn2+, Co2+, and Fe2+); Phys. Rev. B., 2006, 73, 205112. 74. Fuentes-Cabrera, M.; Sumpter, B.G.; Sponer, J.E.; Sponer, J.; Petit, L.; Wells, J.C.; Theoretical study on the structure, stability, and electronic properties of the guanine-Zn-cytosine base pair in M-DNA; J. Phys. Chem. B, 2007, 111, 870–879.

Part B Medical Applications

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

5 Platinum Drugs, Nucleotides and DNA: The Role of Interligand Interactions Giovanni Natile and Francesco Cannito

5.1 Introduction Cisplatin, cis-diamminedichloridoplatinum(II), was the first inorganic antitumour drug used in clinics1 and, after more than 30 years, remains one of the most widely used and successful drugs in cancer therapy.2,3 Today cisplatin is routinely used, alone or in combination with other anticancer drugs, for the treatment of lung, ovary, testes and bladder cancers4 and shows activity against slow-growing as well as rapidly growing tumours. However, as with all chemotherapeutic drugs, cisplatin also has drawbacks, including intrinsic or acquired resistance and toxicity. Toxicity and resistance both have a negative influence on the cure rate.5,6 Some of the severe safety issues have been solved by the introduction of carboplatin, which is equally effective as cisplatin, but with a more acceptable sideeffect profile (Figure 5.1). A key development, in more recent years, has been the elucidation of the mechanism of tumour resistance to cisplatin which includes decreased membrane transport of the drug,7,8 increased cytoplasmic detoxification,9,10 increased DNA repair,11–13 and increased tolerance to DNA damage.14–16 The efforts to circumvent the mechanisms of resistance and thereby broaden the clinical utility of this class of agents have resulted in the discovery of oxaliplatin Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

Platinum Drugs, Nucleotides and DNA

136

H3N

H3N

Cl

Cl Pt

Pt H3N

Cl

Cl

NH3

cisplatin

transplatin

Pt H3N

H2 C

H2 C

O C

O

H3N

C O

CH2

C

C H H2 C

C H N H2

C H2

C H2

O

H2 C

carboplatin

O

H2 N

C

O Pt C

O

O

oxaliplatin O

H3 C C O

Cl

H3N CH2

H C CH2 CH2 CH2

H C

Pt N H2

Cl

Pt N

Cl

O HC

CH2

H3C

C C H

C

satraplatin

HC

Cl

H3N

CH3

O

picoplatin

Figure 5.1 Structures of cisplatin and transplatin, and of other platinum drugs already approved or in the pipeline

(active in patients with colorectal cancer), satraplatin (the first orally administered platinum drug, which shows promise in patients with prostate cancer) and picoplatin (currently in phase III trials against small-cell lung cancer, prostate cancer and colorectal cancer). These drugs are also shown in Figure 5.1. Improved delivery of platinum drugs to tumours is also being studied using liposomal-based17,18 or copolymer-based products,19–21 localized administration22 and also coadministration with specific modulators to prevent platinum-resistance mechanisms.23–26 Reference 27 gives a very recent overview of the field. Cisplatin is a simple neutral inorganic compound having square planar geometry and containing a platinum(II) centre bonded to two nonlabile ammine ligands (carrier ligands) and two labile chlorido ligands (leaving groups) in a cis configuration. Transplatin has the same composition but the ligands are in a trans arrangement. Whereas cisplatin is active against a wide variety of tumours, transplatin is inactive and toxic.28 The structures of cisplatin and transplatin are also shown in Figure 5.1. Since cisplatin and its analogues were found to be tumour active, while transplatin and its analogues were found to be inactive, it was suggested that, in order to

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137

be tumour active, platinum compounds need to satisfy specific requirements, which are known as classical structure–activity relationships.29

5.1.1 Structure–Activity Relationships (SAR) The ligand exchange kinetics of Pt compounds appear to have a great influence on the activity of these drugs.30 Complexes with firmly coordinated leaving groups do not have antitumour activity; also very labile leaving groups do not lead to high activity.31,32 Thus inactivity can be caused by either a too low or a too high reactivity.30 In the latter case the inactivity could be a consequence of rapid reaction with other intracellular components generally referred to as platinophiles. Coordination of a platinophile exerting a high trans effect can also cause release of the otherwise nonexchangeable am(m)ine ligands. The nature of the nonleaving groups also influences the activity of platinum compounds. Several compounds of general formula cis-[PtCl2(amine)2] with different amine ligands have shown antitumour activity. These can be monodentate amines (such as NH3) or bidentate amines like ethylenediamine (en) or 1,2diaminocyclohexane (dach). The activity of the platinum complexes decreases along the series NH3 > RNH2 > R2NH > R3N (R is an alkyl substituent),33 thus indicating that the steric hindrance and the hydrogen-bonding ability of the ligands can be important factors in determining the activity.34 The amines can act as hydrogen donors toward the O6 atom of a guanine or the phosphate groups of DNA.35 These interactions can be important from both a thermodynamic (stabilization of the Pt– DNA adduct) and a kinetic point of view (delivery of the Pt complex to the N7 of guanine). All these observations have resulted in a list of structural requirements for a platinum complex to be endowed with antitumour activity, the so-called structure–activity relationships (SAR):29 (1) Complexes should be neutral. (2) A cis geometry is required with general formula cis-[PtX2(amine)2] for platinum(II), and cis, trans, cis-[PtX2Y2(amine)2] for platinum(IV). (3) The X ligands (leaving groups) should be of intermediate strength (such as Cl−, SO2− 4 or carboxylate ligands). For platinum(IV) complexes the Y ligands should have trans orientation and can be Cl−, OH− or [OC(O)CnH2n+1]−. (4) The nonleaving amine ligands should contain at least one N-bound proton, necessary for hydrogen-bonding interactions with DNA (e.g. H-bonding to O6 of guanines or 5′ phosphate groups). All second and third generation platinum drugs (carboplatin, oxaliplatin) and those in the pipeline (satraplatin, picoplatin) satisfy the SAR given above. However it has been found that several series of platinum compounds which violate at least one of the above described structure–activity relationships are also antitumour active.36 For example, there are compounds with trans geometry37 or containing multiple metal centres that are antitumour active.38 Because of a different mode of interaction with DNA, such compounds are expected to exhibit a completely different spectrum of activity.

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5.1.2 DNA Binding Because platinum belongs to the group of ‘soft’ metal ions, it reacts preferentially with N atoms rather than O atoms. In fact, the preferred binding site in DNA is N7 of purine nucleobases.39–45 It has been suggested that in vivo the positively charged cis-[Pt(NH3)2(OH2)2]2+ ion diffuses to the negatively charged DNA and then rapidly migrates along the helix to the preferred d(GpG) binding sites.46 A scheme with all potential binding sites is represented in Figure 5.2. In double-stranded DNA the only nitrogen atoms which are not involved in Watson–Crick hydrogen bonds and are available for coordination to Pt are N3 and N7 of purines. However N3 is sterically hindered, therefore only N7 is left (bold arrows in Figure 5.2). In single-stranded DNA N1 of adenine and N3 of cytosine are also capable of Pt binding (dotted arrows in Figure 5.2). Finally, all protonated nitrogen atoms of purines and pyrimidines have the lone pair of electrons delocalized on the aromatic p-system of the ring they belong or are attached to, therefore they become available for coordination to Pt only after deprotonation (empty arrows in Figure 5.2).

NH2

5 6 HO

4

3

cytosine (C)

N

2

1 N

O

O H

O H

H

H O

O

guanine (G) N 7 8 9 N

H

P

O

6 5 4

3 N

O H

O–

N 7 5 8 9 4 N

H O

H O

P O

NH2 NH 2

H

H

O

1 NH 2

O

P

adenine (A)

3 N O

5

H O

= preferential platination sites in double-strand DNA

N

H

H

= potential metal binding sites only after loss of a proton

1 2

O H

–

= additional metal binding sites in single-strand DNA

6

H

6

O

O

O–

H

H

OH

H

H

4 1 N

3 NH 2 O thymine (T)

H

Figure 5.2 Possible platinum binding sites on DNA

Introduction

139

N7 of both adenine and guanine can be platinated, however N7 of guanine shows a greater kinetic preference.35,42 This tendency results from the stronger basicity of that nitrogen and from possible simultaneous hydrogen-bond interactions between am(m)ine protons of cisplatin and O6 of guanine.34 In contrast, in the case of adenine, only repulsive interactions can be produced between the NH2 in position 6 of adenine and a platinum-bonded amine ligand.47 The reaction of cisplatin with DNA leads to the formation of at least six different types of Pt–DNA adducts, the most important being schematically depicted in Figure 5.3. The major adduct formed by cisplatin was found to be the 1,2intrastrand crosslink between adjacent guanine bases (d(GpG), 60–65%), followed by the 1,2-intrastrand crosslink between adjacent adenine and guanine bases (d(ApG), 20–25%); 1,3 and 1,4-intrastrand crosslinks between purines separated by one (d(GXG), X stands for a deoxyribonucleotide, 5–6%) or two intervening bases (d(GXXG)) can also occurr.48–51 It can be noticed how intrastrand d(GpG) and d(ApG) adducts together account for about 90% of the total. A small percentage of cisplatin was found to be involved in interstrand crosslinks between two guanines on opposite strands of the DNA, in monofunctional adducts in which the platinum

NH3 (b)

Pt NH3

NH3

Pt

(c) NH3

(d) NH3 NH3

(a)

Pt

NH3

NH3 Pt

PROTEIN

Figure 5.3 Main adducts formed in the interaction of cisplatin with DNA: (a) 1,2-intrastrand crosslink; (b) 1,3-intrastrand crosslink; (c) interstrand crosslink; (d) protein–DNA crosslink

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moiety is coordinated to a single purine, and in DNA/protein crosslinks in which cisplatin coordinates a nucleobase and a protein molecule.49,52–56 The structure of platinated DNA is significantly distorted, resulting in a decrease in melting temperature,57 shortening,58 unwinding59 and local denaturation.60 Results obtained in numerous cell lines suggest that cisplatin-damaged DNA causes cell cycle perturbation and arrest in the G2-phase to allow for damage repair. In the case of inadequate repair, the cell eventually undergoes an abortive attempt at mitosis that results in cell death via an apoptotic mechanism.56,61–64 Transplatin is not able to form 1,2-intrastrand crosslinks because of the trans disposition of the two coordination sites; instead it may form 1,3-intrastrand crosslinks between two G residues separated by at least one base. The amount of monofunctional adducts and protein–DNA crosslinks is much higher for transplatin than for cisplatin. Another difference between the two isomers is that interstrand crosslinks formed by transplatin are between complementary G and C residues, whereas cisplatin forms only interstrand crosslinks between G residues. It was reported that, under physiological conditions, transplatin forms mainly monofunctional adducts, thus allowing the formation of protein–DNA crosslinks.64 In vivo, transplatin interstrand crosslinks might not form because of the slow rate (t1/2 ≈ 24 h) of the crosslinking reaction and the trapping of the monoadduct species by thiolcontaining amino acids.65,66 As stated earlier, DNA binding of platinum complexes is considered to be responsible for their cytotoxic effect. DNA containing platinum adducts is structurally distorted with respect to normal B-DNA, resulting in a loss of stability. The structural aspects of platinated DNA have been thoroughly investigated using NMR spectroscopy,67–71 X-ray crystallography72–74 and gel electrophoresis.75

5.1.3 Model Compounds cis-A2PtG2 complexes, in which A2 is a bidentate amine or two monodentate amines and G is a guanine derivative bound to platinum via N7, are the most simple models of the G/G crosslinks formed by cisplatin.76–83 However, even these most simple models can adopt conformations differing in the relative orientation of the two guanine bases. The G bases can be in a head-to-head arrangement (HH), in which both H8 atoms lie on the same side of the platinum coordination plane, or in a head-to-tail arrangement (HT), in which the H8 atoms lie on opposite sides of the coordination plane (LHT and DHT, Figure 5.4). In the solid state, the large majority of cis-A2PtG2 complexes adopt the conformation with the six-membered rings of the two guanines on opposite sides of the platinum coordination plane.84–89 In only a few cases (all complexes contained 9ethylguanine except one that contained 9-(2-methoxy-ethoxy)guanine) was the HH conformation found.85,90,91 In solution, cis-A2PtG2 complexes usually exhibit free rotation about the Pt–N7 bonds.92–95 In a pioneering study, Cramer demonstrated that guanosine rotation can be slowed sufficiently to detect atropisomers on the NMR timescale, if A2 is a sufficiently bulky chelate (e.g. N,N,N′,N′-tetramethylethylenediamine, Me4en).92 Two G

Introduction

N

N

N

N

N

141

N

Pt

Pt

Pt

∆HT

HH

ΛHT

Figure 5.4 DHT, HH, and ΛHT conformers. The G base is represented by an arrow, the tip indicating H8. The chiralities of the two HT conformers are defined according to the handedness of two straight lines: one perpendicular to the coordination plane and passing through the platinum atom and the second connecting the O6 atoms of the two Gs

H8 signals were detected. In an insightful analysis, it was shown that, when the cisA2Pt moiety has local C2 symmetry, the asymmetric sugar renders the two possible HT atropisomers magnetically nonequivalent, each with one H8 signal. Two H8 signals are also expected for the nonequivalent Gs of the HH atropisomer. The observation of only two of the four possible signals could be best explained by the presence of only the two HT atropisomers, consistent with most crystallographic results.84–89,96–99 The bulky A2 ligand(s) often lacked NH groups. Although N,N′dimethylethylenediamine (Me2en) complexes also formed HT atropisomers, the broad guanosine H8 signals revealed a rotation rate that was moderately fast on the NMR timescale.94 In conclusion, there is an intrinsic difficulty in investigating dynamic nucleos(t)ide complexes. The structure in the solid state may be very different from that in solution because of crystal-packing interactions. In solution, because of fast interconversion between possible conformers, only one set of signals, the average of those of individual conformers, is observed. 5.1.4 Retro Modelling The ‘dynamic motion problem’ led us to construct analogues of cisplatin with bulky ligands designed to reduce the dynamic motion by destabilizing the transition state for Pt-N7 rotation (Figure 5.5). An important feature of the design was to minimize steric effects in the ground state equilibrium species to allow conformers likely to be present in dynamic cis-A2PtG2 adducts to exist in the new adducts also. We introduced the term ‘retro modeling’76 to emphasize that the models we employed76–81,100–105 are more complicated than the relevant molecule, cisplatin. By reducing rotation rates by a billion fold,76,77 retro models have enabled us to understand the adducts of the highly fluxional cisplatin drug with DNA constituents.105 Retro model results76–78,80,81,100,101,103,104,106 have called into question two concepts that for many years were widely accepted from studies on dynamic platinum complexes: (i) untethered Gs adopt an HT conformation in preference to the HH conformation107 and (ii) single-stranded d(GpG) crosslinks favour the HH form, which undergoes slow rotation about the Pt-N7 bonds.108–112

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Platinum Drugs, Nucleotides and DNA l Chelate-ring puckering

d Chelate-ring puckering

H

H

Me

Me N H

N Me

Pt

(S,R,R,S)-Me2dab-Pt

Me

Pt

H

(R,S,S,R)-Me2dab-Pt

H N

N

N

H

N

N

N

Pt

Pt

(S,R,R,S)-bip-Pt

(R,S,S,R)-bip-Pt

H

H

H

H H

N H

N Pt

N

N H

Pt

(S,R)-pipen-Pt

H

(R,S)-pipen-Pt

Me

Me

Me

Me N

Me

N Pt

N

N Me

Me

(R,R)-Me4dach-Pt Me

Pt

Me

(S,S)-Me4dach-Pt Me

Me

Me N

Me

N Pt

N

N Me

(R,R)-Me4dab-Pt

Me

Pt

Me

(S,S)-Me4dab-Pt

Figure 5.5 Me2dab, bip, pipen, Me4dach, and Me4dab ligands with in-plane bulk used to slow-down rotation about the Pt-N7 bonds in cis-A2PtG2 compounds

N,N′-dimethyl-2,3-diaminobutane (Me2dab) was the first carrier ligand used in such a detailed NMR retro model study.78,82,83 Me2dab differs from Me2en in having an additional methyl substituent on each carbon atom of the ethylene chain bridging the two nitrogens. These two methyl substituents not only increase the overall steric rigidity of the ligand, but also, by asymmetric induction, influence the configuration of the adjacent amine groups. Each N and each C in the ring is a centre of asymmetry. The Me2dab retro models provided the opportunity to define the solution structure of conformers by NMR methods, allowing us to identify the HH form of a cis-A2PtG2 adduct for the first time105 and also allowing us to use the known

Introduction

143

Me2dab configuration to define the absolute conformation of the HT forms in solution for the first time.78 The 2,2′-bipiperidine ligand (bip) is analogous to Me2dab, but is much more sterically rigid. This rigidity has two components: first, the chelate ring is part of a three-ring system, decreasing its fluxional character with respect to the Me2dab ligand; second, the CH groups projecting toward the G coordination sites are unable to rotate away from the G bases during G base rotation around the Pt-N7 bond. The bip ligand was able to decrease the dynamic motion roughly one billion times relative to (NH3)2 and ca. 100 times relative to Me2dab in cis-A2PtG2 complexes.76–78,101,104,106,113 Most of our work involved the C2-symmetrical isomers of Me2dab and bip, in which the chiral N, C, C and N centres in the chelate-ring had either S,R,R,S or R,S,S,R configurations. 5.1.5 Retro Models with Untethered Guanine Bases The use of Me2dab and bip ligands allowed the simultaneous observation by 1H NMR spectroscopy of all possible conformers (two HT and one HH conformer, Figure 5.6) that can be formed in cis-A2PtG2 complexes with a C2-symmetrical carrier ligand. Moreover, the conformers were present at equilibrium in different amounts depending upon the chiralities of the carrier ligands; thus these ligands were named chirality-controlling chelates (CCC). The relative amounts were the same for bip and Me2dab compounds with a given chirality and they differ only in dynamics.

Free 5'–GMP HHd

9.0

HTd

8.7

8.4

HHu

HTu

8.1

ppm

Figure 5.6 H8 region of the 1H NMR spectrum for (R,S,S,R)-Me2dab-Pt(5′-GMP)2 (Ref. 53 for related work)

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The equivalent bases in each HT form had H8 chemical shifts intermediate between those of the nonequivalent bases in the HH form. Moreover the H8 signal of the major HT form appeared downfield from the H8 signal of the minor HT form.76–78,82,104 The H8 chemical shift is influenced by the positioning of the H8 atom with respect to the shielding cone of the cis G. The canting of the base in the direction that moves H8 away from the cis G will lead to less H8 shielding and hence to a downfield H8 signal relative to the average H8 signal (this relationship is called ‘6-in’ because the six-membered ring of G moves closer to the cis G). In contrast, a rotation of the base in the direction moving H8 toward the cis G will lead to greater H8 shielding,110 and hence to a more upfield H8 signal (this relationship is called ‘6-out’ because the six-membered ring of G moves further from the cis G, Figure 5.7). The G base of the HH form with O6 on the same side of the coordination plane as the N-H of the cis amine had an upfield H8 signal (HHu), indicative of an H8

S,R,R,S chirality of Me2dab or bip P

P

P O

N

N Pt N

N Pt N

P

P

O

O

O

Pt N H O

O P

ΛHT (major)

∆HT (minor)

HH

R,S,S,R chirality of Me2dab or bip P P

O

P

O

P N Pt N O

N Pt

N

O

N Pt

P

O

N O

P ΛHT (minor)

∆HT (major)

HH

Figure 5.7 HT and HH conformers in Me2dab or bip-Pt(5′-GMP)2 complexes having (S,R,R,S) or (R,S,S,R) chirality at the carrier ligand. N-CH2X groups (X = H and R for Me2dab and bip, respectively) are indicated by larger circles, N-H protons by smaller circles. Only the chelate ring atoms of the carrier ligands are shown. 5′-GMP is represented by an arrow, the tip indicating H8 and the tethered P the sugar-phosphate group

Internucleotide Interactions 145

pointing toward the cis G (‘6-out’ canting). The base with O6 on the same side of the coordination plane as the N-CH2X group of the cis amine (X = H or R for Me2dab and bip ligands, respectively) had a downfield H8 signal (HHd),76–78,82,104 indicative of an H8 pointing away from the cis G (no canting or ‘6-in’ canting). Differently from HH, in HT conformers the two bases are equivalent and the H8 of each G is on the same side of the coordination plane as the six-membered ring of the cis G. Under these circumstances the shielding effect of the cis G is expected to be significant even for no canting or moderate ‘6-in’ canting of the two nucleobases.

5.2 Internucleotide Interactions 5.2.1 Dipole–Dipole Interaction Between cis Guanines The major HT conformer found in Me2dab and bip-PtG2 adducts has G O6 located on the opposite side of the coordination plane from the cis amine N-H.78,104 The favoured form could not participate in a G O6/NH cis amine hydrogen bond, revealing that such an H-bonding might not be so important (Figure 5.7).100,104,114 The most plausible explanation for this behaviour is that stabilization of the HT conformers stems from dipole–dipole interactions between cis bases, and that this interaction is stronger for G bases having the six-membered ring of each guanine leaning towards the cis G (‘6-in’ canting). The ‘6-in’ canting, however, will bring the H8 of each guanine close to the cis amine and, if an N-Me substituent is on the same side of the platinum coordination plane as the H8, an unfavourable interaction between the two groups will limit the degree of ‘6-in’ canting and the stability of the conformer. This is the case for ‘minor’ HT conformers depicted in Figure 5.7. It is noteworthy that, in X-ray structures of HT conformers of cis-A2PtG2 complexes, much greater ‘6-in’ canting can be found when the H8 of each guanine is on the same side of the platinum coordination plane as an N-H of the cis amine115,116 than when it is on the side of an N-Me of the cis amine.117,118 A ‘6-in’ canting of ca. 20 ° (deviation from the guanine plane orthogonal to the coordination plane) will bring the electron-rich O6 atom of each guanine closer to the electron-deficient H8 atom of the cis base. Moreover, greater canting also reduces the dihedral angle between the planes of the two guanines (for a canting angle of 10 °, 20 °, 30 ° and 40 ° the dihedral angle is found to be 82 °, 74 °, 66 ° and 56 °, respectively) and possibly increases internucleotide stacking interactions. 5.2.2 Repulsion Between O6 Groups of cis Guanines In Me2dab and bip-PtG2 HH conformers, there is a large dispersion of H8 chemical shifts (∼1 ppm) that stems from a combination of electronic and steric effects. Electrostatic repulsion between electron-rich O6 atoms of cis Gs will tend to place the six-membered rings farther apart.119 The stereochemistry of the Me2dab and bip

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ligands (for R,S,S,R or S,R,R,S configurations at the N, C, C and N asymmetric centres) places one N-H and one N-CH2X on each side of the platinum coordination plane. The guanine with the six-membered ring on the side of the coordination plane as the N-H of the cis amine can be canted ‘6-out’ because there is no clash with an amine substituent and O6 can act as an H-bond acceptor from the N-H. In contrast, the guanine with the six-membered ring on the side of the N-CH2X group of the cis amine will be much less canted and probably canted away from the cis amine (‘6-in’ canting). The former guanine will have a strongly shielded H8 signal, whereas the second guanine will have an unshielded H8 signal (HH conformers in Figure 5.7). Removal of the electron-rich O6 atom of guanine, such as in the case of adducts with benzimidazole (B), not only increases the yield of the HH conformer with respect to the HT conformers (the dipole–dipole interactions stabilizing the latter conformers are greatly reduced, if not absent), but also greatly reduces the dispersion of the B H2 chemical shifts even in the case of Me2dab or bip ligands (B H2 occupies the same position as H8 in G).119 This behaviour is sketched in Figure 5.8.

5.2.3 Phosphate N1H cis G H-Bond Interaction, SSC Two relevant types of ionizable groups are present in cis-A2Pt(GMP)2 adducts. First, the monoanionic phosphates of the GMP ligands [ROP(OH)O2]− which have pKa values (5.6 to 6.2)84–86,112 lower than for free 5′-GMP (6.3)76,79,81-83 and become fully deprotonated by pH ∼7.5.77–80,84 Second, the N1H of each G which deprotonate with pKa values of 8.7 and ∼9.479,83,87,109 versus 9.5 for free 5′-GMP.79,80,84 For both the R,S,S,R and S,R,R,S chiralities of the Me2dab and bip-PtG2 complexes, the percentage of the DHT conformer in the case of 3′-GMP and the percentage of the LHT conformer in the case of 5′-GMP were higher near pH 7 than at lower pH.76,79–81,102,105 We have unravelled the reasons for this difference in the preferred conformer.104 The phosphate group position is different in 5′-GMP versus 3′-GMP. For 5′-GMP adducts, the 5′-phosphate group projects toward the cis nucleotide in the LHT form, and away from the cis nucleotide in the DHT form (assuming that the nucleotide maintains the preferred anti conformation, Figure 5.9). In contrast, for 3′-GMP adducts the 3′-phosphate is close to the cis nucleotide in the DHT form and away from it in the LHT form (this can be seen in a gedanken experiment in which the phosphates of Figure 5.9 are detached from the 5′ position and attached to the 3′ position of the ribose). Thus, the relative positions of phosphate groups to the cis G in a HT form with a given chirality are opposite for 3′-GMP versus 5′-GMP adducts. On these grounds the key factors stabilizing the DHT conformer for 3′-GMP derivatives and the LHT conformer for 5′-GMP adducts near pH 7 appear to be the phosphate/N1H cis G interactions. This represents a third type of internucleotide interaction. We call such interligand interaction ‘second-to-second sphere communication’ (SSC) because the interacting groups are at the periphery of the cis nucle-

Internucleotide Interactions 147

Figure 5.8 Different ratio of HH and HT conformers in adducts with G (top) and 5,6 dimethylbenzimidazole (bottom). The scheme does not intend to take into account the chirality of the HT conformer (D or Λ) (Ref. 119 for related work)

otides.81,104,105 This is not the case for the previous two types of internucleotide interactions which involve parts of ligands close to the metal and can be referred to as ‘first-to-first sphere communication’ (FFC). SSC interactions are optimal at pH 6–7, conditions in which the phosphate group is deprotonated while N1 is still protonated. Further support for the conclusion that in cis-A2Pt(5′-GMP)2 complexes the LHT conformer is favoured by G phosphate/N1H cis G hydrogen bonding (SSC) came also from our recently reported X-ray crystal structure of Me4en-Pt(5′GMP)2.118 Three unique features not present in all other cis-A2Pt(GMP)2 solid-state structures were found: a LHT conformation, very strong hydrogen-bond interactions between phosphate and N1H of cis nucleotides, and a very small dihedral angle between the planes of the two guanines, although both were nearly perpendicular to the coordination plane.

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platinum

hydrogen

nitrogen

carbonium

oxygen

phosphorus

Figure 5.9 (Plate 7) X-ray structures of two cis-A2Pt(5′-GMP)2 adducts showing how in the DHT conformer (top, based on data from Ref. 117) the 5′-phosphate is directed towards the cis amine, while in the ΛHT conformer (bottom, based on data from Ref. 118) the 5′phosphate is directed towards the cis guanine. For simplicity only the chelate ring atoms of the carrier ligands (Me4dach in the former case and Me4en in the latter case) are shown (See colour plate section)

Guanine cis Amine Interactions 149

5.3 Guanine cis Amine Interactions 5.3.1 Steric Interaction between G and cis Amine As already pointed out in Section 5.2.1, in HT conformers having ‘6-in’ conformation, the relevant steric interactions involve the G H8 and the cis amine. Hence the major HT conformer found for Me2dab and bip-PtG2 adducts has the guanine H8 located on the same side of the coordination plane as the cis amine N-H and on the opposite side from the cis amine N-CH2X group (‘major’ HT conformers in Figure 5.7). In the HH conformer (Section 5.2.2) the guanines tend to have ‘6-out’ conformation and therefore the relevant interactions involve the guanine six-membered rings and the cis amine substituents. In order to have moderately stable HH conformers it is required that at least one guanine has the six-membered ring on the side of the coordination plane where the cis amine has a nonbulky substituent (such as an N-H). Steric clash between the six-membered ring of each guanine and the cis amine substituent positioned on the same side of the platinum coordination plane renders the HH conformer particularly unstable in adducts with tertiary amines (such as Me4dab and Me4dach reported in Figure 5.5)117,118,120 or N-donor heterocycles (such as phenanthroline and 2,9-dimethylphenanthroline).121 5.3.2 Guanine O6-NH cis Amine H-Bond Interactions At basic pH (∼10), the N1H deprotonation eliminates the SSC phosphate/N1H cis G interactions that stabilize the LHT conformer for 5′-GMP adducts and the DHT conformer for 3′-GMP adducts (Section 5.2.3). On the other hand, N1H deprotonation greatly increases the capacity of the G O6 to act as a hydrogen bond acceptor. It was observed that, upon N1H deprotonation at basic pH, the less abundant HT conformer of the moderately dynamic Me2dab-PtG2 model compounds becomes more favoured (Table 5.1). (Such an increase in the concentration of the minor HT conformer at basic pH is not observed for the R,S,S,R configuration of the diamine and 5′-GMP nucleotide for the reason discussed in the next section.)113 The ‘minor’ HTs have the G O6 of each guanine on the same side of the platinum coordination plane as the N-H of the cis amine (Figure 5.7); therefore, their increase in concentration at basic pH is likely to stem from G O6/NH Me2dab hydrogen bonding, now favoured as a result of an increased capacity of the G O6 to act as a hydrogen bond acceptor. 5.3.3 Phosphate/NH cis Amine H-Bond Interaction, FSC When G has a 5′-phosphate group, conformer distribution could also be influenced by phosphate/NH cis amine hydrogen-bond interactions. Such an interaction should favour the DHT conformer because for this conformer the 5′-phosphate protrudes toward the cis amine (assuming the nucleotides have the favoured anti conformation, Figure 5.7). As noted in the previous section, at basic pH the LHT conformer is unexpectedly disfavoured (Table 5.1) for (R,S,S,R)-Me2dab-Pt(5′-GMP)2 although

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Table 5.1 Conformer percentages of Me2dab-PtG2 complexes at different pH values measured from the H8 signal intensities. Increase of pH from ∼3 to ∼7 highlights the effect of SSC (stabilization of DHT in the case of 3′-GMP adducts and stabilization of ΛHT in the case of 5′-GMP adducts), while further increase of pH from ∼7 to ∼10 highlights the effect of guanine O6/NH cis amine H-bond interaction (stabilization of DHT in the case of S,R,R,S configuration of the diamine and stabilization of ΛHT in the case of R,S,S,R configuration of the diamine). The latter stabilization (ΛHT favoured at high pH for R,S,S,R configuration of the diamine) is not observed in the case of 5′-GMP because of simultaneous stabilization of the DHT conformer stemming from 5′-phosphate/NH cis amine H-bond interaction as outlined in Section 5.3.3. (S,R,R,S) G 3′-GMP 5′-GMP

(R,S,S,R)

pH

% LHT

% DHT

% HH

pH

% LHT

% DHT

% HH

3.4a 7.2a 9.6a 3.2 7.0 10.4

80 60 37 92 98 20

13 35 60 2 1 72

7 5 3 6 1 8

3.3 6.9 9.3 3.1 7.3 10.0

16 4 86 7 12 10c

84 96 14 71 70 69

b b b

22 18 21

a

At 5 °C; all others were at room temperature. Not determined. The percentage of this conformer (ΛHT) does not increase at the expenses of the other HT conformer (DHT) when the pH is raised to 10. b c

the G O6 and the N-H of the cis amine are on the same side of the platinum coordination plane (Figure 5.7). We believe that the nonincrease of the LHT concentration at basic pH is due to a contemporary increase in stability of the DHT conformer with the consequence that, on moving from neutral to basic pH, the conformer concentrations remain practically unchanged. The stabilization at basic pH of the DHT conformer most probably stems from a phosphate/NH cis amine interaction (as depicted in Figure 5.7). Indeed, for the DHT conformer of (R,S,S,R)-Me2dabPt(5′-GMP)2 we match the situation in which the 5′-phosphate protruding towards the cis amine is on the same side of the platinum coordination plane as the N-H of the cis amine. In Me2dab-Pt(3′-GMP)2 adducts, conformers cannot be affected by phosphate/NH cis amine interaction for stereochemical reasons because the 3′phosphate is too remote from the cis amine N-H. Because the 5′-phosphate/NH cis amine hydrogen bonding involves one part of the ligand close to the metal (the NH) and one part far from the metal (the phosphate), we have called this interaction ‘first-to-second sphere communication’ (FSC). This is not the case for the previous two types of guanine/cis amine interactions which involve parts of ligands close to the metal and can be referred to as ‘first-to-first sphere communication’ (FFC).

5.4 Solid-State Structures of Dynamic Nucleotides In the solid state, numerous examples of DHT complexes of 5′-GMP and several related 5′-phosphate derivatives have been found, not only for Pt with various

Conformer Distribution in Cisplatin Adducts 151

carrier ligands, but also for other metals.85–93,96,110,113,115 The nucleotides in the structures have very similar relationships, with the purine bases having the same relative positions and the phosphate groups always hydrogen-bonded to the cis ligands in a similar manner. Only very recently has a LHT complex of 5′-GMP been reported,118 while no example of a cis bisnucleotide HH complex has yet been published. This contrast between the prevalence of FSC in the solid and its relative unimportance in solution supports the premise of our retro model studies that the structures in the solid state of dynamic nucleotide complexes may be very different from the solution structures. Interligand interactions, as they result from our retro model studies, can be grouped in two distinct categories: nucleotide/nucleotide and amine/nucleotide. Internucleotide interactions concern: (1) Electrostatic attraction in HT conformers between electron-rich guanine O6 and electron-deficient H8 of cis Gs, both on the same side of the platinum coordination plane. This interaction is responsible for these conformers generally having the six-membered ring of each guanine leaning towards the cis G so to bring the two oppositely charged moieties closer to each other (‘6-in’ conformation). (2) Electrostatic repulsion in the HH conformer between electron-rich O6s, both located on the same side of the platinum coordination plane. This interaction generally leads to ‘6-out’ canting of the guanines. (3) H-bond interaction between nucleotide phosphate and N1H of cis G (SSC). In the case of 5′-phosphate this interaction will favour LHT (since in LHT each anti 5′-GMP has the 5′-phosphate protruding towards the cis G), while in the case of 3′-phosphate will favour DHT (since in DHT each anti 3′- GMP has the 3′-phosphate protruding towards the cis G). Amine/nucleotide interactions can also be of different types: (1) Steric interaction between amine and cis G. In HT conformers (having preferentially the ‘6-in’ conformation) the relevant interaction involves the guanine H8 and the cis amine substituents, while in the HH conformer (having preferentially the ‘6-out’ conformation) the relevant interaction involves the sixmembered ring of the guanine and the cis amine substituents. (2) H-bond interaction between guanine O6 and N-H of the cis amine (when present). This interaction becomes significant after deprotonation of guanine N1H at basic pH. (3) H-bond interaction between nucleotide 5′-phosphate and N-H of the cis amine when present (FSC). For HT conformers, assuming an anti conformation of 5′GMP, only D chirality allows for such an interaction.

5.5 Conformer Distribution in Cisplatin Adducts of G Derivatives NMR methods do not allow detection of individual conformers in fluxional cisA2PtG2 complexes with two ammines or a primary diamine ligand. Because of this dynamic motion problem, we relied heavily on CD spectroscopy to assess the chirality of the major HT conformer.

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5.5.1 CD Signature of Conformers The presence of enhanced CD signals for some cis-A2PtG2 complexes was first reported in 1980,122 and attempts were later made to interpret the results on a structural basis.123,124 However, a convincing interpretation of the results has depended on subsequent studies with less dynamic complexes.77,78,107 The CD spectra of (S,R,R,S)-Me2dab-PtG2 complexes all had the same shape, i.e., positive peaks at ∼285 and 230 nm and negative peaks at ∼250 and 210 nm. Because all the 2D NMR studies showed that the dominant atropisomer was LHT (Figure 5.7), this type of CD signal was designated as L. The CD spectra of enantiomeric (R,S,S,R)-Me2dab-PtG2 complexes had signs opposite to those of the corresponding peaks in the (S,R,R,S)-Me2dab-PtG2 CD spectra (negative peaks at ∼285 and 230 nm and positive peaks at ∼250 and 210 nm). Because the 2D NMR studies showed that the dominant atropisomer for (R,S,S,R)-Me2dab-PtG2 complexes had the DHT conformation (Figure 5.7), the latter type of CD signal was designated as D. Furthermore, the CD signal intensity roughly correlated with the percentage of the major HT form.78 Studies with the related bip complexes indicated that a mixture of 25% DHT, 50% HH, and 25% LHT had essentially no CD intensity.77 The CD intensity increased as the favoured HT form became dominant with time. Similarly, in a ‘pH jump’ experiment performed on (S,R)-pipen-Pt(5′-GMP)2 (Figure 5.5) in which we monitored the change of atropisomer distribution by using both NMR and CD spectroscopy,79 changes in H8 NMR signal intensities corresponded to changes in CD intensity. On the basis of the above experiments, we interpret the sign of the CD signal as a reflection of the D or L conformation of the major HT form. In particular, the pattern of a positive Cotton effect around 285 nm and a negative Cotton effect around 250 nm is characteristic of a LHT conformer; the opposite pattern is characteristic of a DHT conformer.76

5.5.2 Conformers in Cisplatin Adducts with 3¢-GMP: The Role of SSC The CD signals of cisplatin adducts of G derivatives with no phosphate groups are weak. No conformer is favoured because the absence of phosphate groups rules out both FSC and SSC interactions; moreover the two ammine ligands have no chiral property that can exert a stereochemical control through FFC interactions so favouring a particular conformation. This analysis applies not only to cisplatin, but also to analogous A2PtG2 complexes with achiral primary diamines because these also have weak CD signals. In contrast, at neutral pH, cis-(NH3)2Pt(3′-GMP)2, pn-Pt(3′-GMP)2 (pn = 1,3diaminopropane), and en-Pt(3′-GMP)2 all showed the D-type CD signal,105 indicating that the major atropisomer is DHT (Figure 5.10). The carrier ligand has a modulating effect on the CD intensity: weakest for en-Pt, strongest for cis-(NH3)2Pt. The 3′-phosphate group from each 3′-GMP can form a hydrogen bond with the N1H of the cis 3′-GMP in the DHT conformation (assuming the nucleotides keep the normally preferred anti conformation), whereas such hydrogen bonding is less

Conformer Distribution in Cisplatin Adducts 153 a b

∆c (M–1 cm–1)

5

c

a) (NH3)2Pt(3'–GMP)2 b) pnPt(3'–GMP)2 c) enPt(3'–GMP)2 pH = 7

0

–5

250

nm

a) (NH3)2Pt(5'–GMP)2

8

a b) pnPt(5'–GMP)2 b

4 ∆c (M–1 cm–1)

300

c) enPt(5'–GMP)2 pH = 7

c 0

–4

–8 250

300

nm

a) enPt(1–Me–5' GMP)2 b) pnPt(1–Me–5' GMP)2 pH = 7.8

a 6 4 ∆c (M–1 cm–1)

b 2 0 –2 –4 –6 250

300

nm

Figure 5.10 CD spectra of cis-A2PtG2 compounds when A2 = (NH3)2, pn, or en and G = 3′GMP, 5′-GMP, or 1-Me-5′-GMP (Ref. 105 for related work)

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favourable in the LHT conformation. The preference for the DHT atropisomer for these three 3′-GMP complexes appears to be due to such a hydrogen bonding (SSC). Raising the pH from 3 to 7 increased the D type CD signal intensity, indicating that the DHT form is favoured upon 3′-phosphate group deprotonation, most likely due to formation of stronger phosphate/N1H cis G hydrogen bonds at neutral pH. Above pH 7 the D type CD signal starts to decrease, and the intensity is nearly zero at pH 9. At pH ∼9, where N1H deprotonation occurs,125 the N1 atom has lost its proton and no phosphate/N1H hydrogen bond can exist. Our NMR/CD studies with pipen, Me2dab and bip-Pt(3′-GMP)2 complexes support this interpretation.76,102

5.5.3 Conformers in Cisplatin Adducts with 5¢-GMP: Interplay between SSC and FSC At neutral pH, the CD signals of cis-(NH3)2Pt(5′-GMP)2, pn-Pt(5′-GMP)2 and enPt(5′-GMP)2 are of the L type, indicating that the major conformation is LHT (Figure 5.10).105 Intramolecular hydrogen bonding between the phosphate group of one 5′-GMP and the N1H of the cis 5′-GMP (SSC) is more favourable in the LHT than in the DHT conformer. The CD data point to substantial differences in signal intensity among 5′-GMP adducts of different carrier ligands [cis-(NH3)2, pn, en]. One obvious potential factor is the bite angle of the carrier ligand(s). The very low CD intensity of the en-Pt(5′GMP)2 complex suggests that the low bite angle of the en ligand reduces the strength of the SSC interaction. Raising the pH from ∼3 to 7 increased the L type CD signal intensity, indicating that the LHT form is more favoured upon 5′-phosphate group deprotonation, possibly because of stronger phosphate/N1H hydrogen bond at neutral pH. As the pH was raised further, the CD signals decreased in intensity and partially inverted at pH 10, suggesting a bias toward the DHT form after N1H deprotonation.105 The preference for the LHT atropisomer up to pH 9 is likely to stem from the 5′phosphate group of each 5′-GMP forming a hydrogen bond with the N1H of the cis 5′-GMP (SSC). In the LHT conformation the 5′-phosphate protrudes toward the cis nucleotide (assuming the nucleotides keep the preferred anti conformation). On the other hand, the bias towards the DHT form observed at pH 10 can be explained as follows. In the DHT conformation the 5′-phosphate is directed towards and can directly reach the cis amine and form an H-bond (FSC). It is worth nothing that the latter hydrogen bonding interaction is found in relevant X-ray structures. It is therefore most likely that the presence of 5′-phosphate/NH cis amine hydrogen bonding is responsible for the stabilization of the DHT form at high pH (such hydrogen bonding is masked at acidic and neutral pH by the dominating SSC interactions). 5.5.4 Conformers in Cisplatin Adducts with 1-Me-5′-GMP: The Role of FSC The CD signals of pn-Pt(1-Me-5′-GMP)2 and en-Pt(1-Me-5′-GMP)2 had signs opposite to those of 5′-GMP adducts (Figure 5.10).105 The signs indicate that the major atropisomer of the 1-Me-5′-GMP adducts is DHT.

Conformer Distribution in Cisplatin Adducts 155

Because no phosphate/N1H hydrogen bond is possible for 1-Me-5′-GMP derivatives (SSC), the only important interaction influencing the HT atropisomer population is phosphate/NH cis amine H-bonding (FSC). The DHT conformer has the 5′-phosphate protruding toward the cis amine and able to form a phosphate/NH cis amine H-bond. This conclusion is also supported by the observation that in the absence of a cis G (which rules out any SSC interaction), such as in the complex Me3dien-Pt(5′GMP) (Me3dien = N,N′,N′′-trimethyldiethylenetriamine, with all three methyl groups on the same side of the platinum coordination plane), H-bond formation between the phosphate and the N-H of the cis amine becomes the dominant interaction.126,127 5.5.5 Conformers in Oxaliplatin Adducts with Guanines Lacking a Phosphate Group: The Role of FFC In an early investigation of the reaction of enantiomeric dach platinum complexes with guanine derivatives,123 Pasini and coworkers observed that the CD spectrum of (R,R)-dach-PtG2 (G = 9-methylguanine) was characterized by a positive Cotton effect centred at 230 and 280 nm and a negative Cotton effect centred at 260 nm; the corresponding Cotton effects for the (S,S)-dach-PtG2 enantiomer had the opposite sign (Figure 5.11). The Cotton effect was assigned to coupling between p–p* electronic transitions centred on the guanine bases and was taken as a clear indication of transmission of chirality from the dach ligand to the coordinated cis guanines mediated by the amine protons.123 The details of the mode of transmission of chirality given at that time were not convincing and have remained obscure until very recently.128 cis-A2PtG2 complexes, with G = a guanine derivative lacking a phosphate substituent, can have only FFC interactions. However, whereas each N in retro-model carrier ligands had substituents of very different bulk on the two sides of the

(R,R)DACH-Pt(9–EtG)2 (S,S)DACH-Pt(9–EtG)2

∆ε/ (mol–1 cm–1)

2 1 0

200

250

300

λ / nm

350

Figure 5.11 CD spectra of (R,R)-dach-Pt(9-EtG)2 (broken line) and (S,S)-dach-Pt(9-EtG)2 (solid line) at pH 7 and room temperature

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platinum coordination plane, the dach ligand has an N-H on each side of the plane. Nevertheless, a slight difference in the stereochemistry of the two protons (one N-H has ‘quasi axial’ and the other ‘quasi equatorial’ character) is sufficient to induce a significant change in the relative stabilities of the dach-PtG2 DHT and LHT conformers. At acidic and neutral pH the stability of HT conformers is governed by G to G dipole–dipole interaction, which is greater for the six-membered ring of each guanine leaning toward the cis G. Such a ‘6-in’ canting of the two guanines can be hampered by the steric interaction between the H8 of each guanine and the substituent on the cis amine that is on the same side of the platinum coordination plane. Such a repulsion is greater for a ‘quasi equatorial’ N-H than for a ‘quasi axial’ N-H. Therefore the DHT conformer is stabilized in the (S,S)-dach-PtG2 complex, while the LHT conformer is stabilized in the (R,R)-dach-PtG2 species (this can be seen in Figure 5.7 if the larger circles now represent ‘quasi equatorial’ protons and the smaller circles ‘quasi axial’ protons). Indeed the CD spectrum is typical of a DHT conformer in the former case and of a LHT conformer in the latter case.128 At basic pH, deprotonation of the guanine N1-H renders the O6 a much better H-bond acceptor; therefore, the stability of the HT conformers is governed by the guanine O6/NH of the cis amine H-bond interaction. Such a guanine O6/NH cis amine interaction is more favourable for a ‘quasi axial’ than for a ‘quasi equatorial’ N-H. As a consequence, at basic pH the CD spectrum of (S,S)-dach-PtG2 has the signature of a LHT conformer, while the CD spectrum of (R,R)-dach-PtG2 has the signature of a DHT conformer (this can also be seen in Figure 5.7, assuming the larger circles to represent ‘quasi equatorial’ protons and the smaller circles ‘quasi axial’ protons).128 5.5.6 Quantifying the Atropisomer Distribution in Adducts of Cisplatin or Analogous A2Pt(5¢-GMP)2 Complexes with Achiral Primary Diamines CD spectroscopy is an empirical method at the current state of theoretical understanding of CD spectral transitions in these types of complexes, therefore it cannot be used for a quantitative estimate of different conformers, for which we have to rely upon NMR data. Very recently, by lowering the temperature down to 238 K, it has been possible to observe different conformers of (cis-1,4-dach)Pt(5′-GMP)2 (1,4-dach = 1,4diaminocyclohexane for which the N–Pt–N bite angle is ≥97 °). The percentages of the HH, LHT and DHT conformers were estimated from intensities of 1H NMR signals to be 33, 57 and 16%, respectively. This is the first case in which the abundance of the three possible rotamers has been determined in A2Pt(5′-GMP)2 adducts with primary diamines.129

5.6 Retro Models Applied to Adducts with Tethered Guanine Bases The cis-(NH3)2Pt(GpG) and cis-(NH3)2Pt(d(GpG)) complexes, the simplest Glinked models of the major cisplatin–DNA adduct, have been characterized by a

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number of techniques, including CD spectroscopy,109,130 1H NMR spectroscopy108,130 and molecular modeling.110 On the basis of the observation that both nucleotides of each complex were anti (Figure 5.12),130 the crosslinks were initially assigned structures in the HH1 conformer class. Studies of retro models have shown that there are three other classes, namely, HH2, DHT1, and LHT2 (Figure 5.13). Members of adjacent classes are separated by ca. 180 ° rotation of one G about its Pt-N7 bond. Variants in nonHH1 conformer classes have been found for bip-Pt(d(GpG)) and bip-Pt(GpG) adducts.101,103,106 Moreover, the bases can have right-handed (R) or left-handed (L) canting (handedness of canting is defined by the following two straight lines: one connecting the platinum-coordinated N7 atoms of the two guanines and the second connecting the N1 and C8 atoms of a given guanine). The base canting is determined by the chirality of the amine: (S,R,R,S)-bip induces left-handed canting (L, upper row of Figure 5.7) while (R,S,S,R)-bip induces right-handed canting (R, lower row of Figure 7) of the guanine bases. (S,R,R,S)-bip-

O N7

N

O H3' H2'

H5'' H4'

NH2

N

H1' O

H2''

O

O

(a) N7

P O

H

H8

OH

H5'

N

H5'

H

N

H8

O

N

O H3' H2'

H5'' H4'

NH2

N

H1' H2''

HO

H H2N

O N

OH

N

H H H

N

O H H HO

H H H

(b)

N

O

H NH2

N

N

OH H H

N

N

O H

H

H

H

H

H

HO

(c)

Figure 5.12 Structure of d(GpG) (a) and schematic anti (b) and syn (c) guanine glycosidic torsion angle alignments. The H8 and sugar H1′ protons are closer to each other (2.5 Å) in the syn alignment compared to a longer distance (3.7 Å) in the anti alignment, a feature that can be readily monitored by NMR

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HH1

∆HT1

HH2

platinum

carbonium

nitrogen oxygen

phosphorus

Figure 5.13 (Plate 8) Minimum energy models (from NMR-restrained MMD calculations) of the anti/anti HH1 L (top) and anti/syn DHT1 L (centre) forms of (S,R,R,S)-bip-Pt(d(GpG)) and the anti/anti HH2 R (bottom) form of (R,S,S,R)-bip-Pt(d(GpG)) (the conformation of the 5′-G is given first, then that of 3′-G; only the chelate-ring atoms of the bip ligand are shown). Based on data from Refs 101 and 106 (See colour plate section)

Pt(d(GpG)) and (S,R,R,S)-bip-Pt(GpG) both favoured two variants, anti/anti HH1 L (with N and S puckers for the 5′ and 3′-G residues, respectively) and anti/syn DHT1 L (both sugars having mainly N puckers, top and centre molecular models in Figure 5.13).103,106 When these complexes were kept at pH 10 for several days, the DHT1 L variant became favoured over the HH1 L variant, demonstrating that DHT1 L is more favourable when N1 is deprotonated.103,131 For (R,S,S,R)-bip-Pt(d(GpG))

Retro Models Applied to Adducts 159

at physiological pH, the two major conformers were anti/anti HH1 R and anti/anti HH2 R (bottom model in Figure 5.13). For both conformers the sugar pucker was N and S for the 5′ and 3′-G, respectively.101 When the sample was kept for 1 day at pH 10, where N1 was not protonated, the LHT2 R conformer became ∼30% abundant.103 In contrast to the derivative with d(GpG), for (R,S,S,R)-bip-Pt(GpG) only the HH1 R variant was observed at any pH, and only a small population of LHT2 R was observed at pH 10.103 5.6.1 Interligand Interactions Stabilizing Different Conformers in Adducts with Tethered Guanines Investigations of adducts of retro models with d(GpG) and GpG dinucleotides, not only revealed the dynamic behaviour of these intrastrand crosslink models, with the possibility for each guanine to undergo ca. 180 ° rotation about the Pt-N7 bond, but also highlighted the role of ‘weak’ interactions, not unlike those discovered in the investigations with untethered cis guanines. In particular, in the HH conformer, a key interaction appeared to be the repulsion between the electron-rich O6 atoms of the two guanines. Because of such repulsions, the six-membered ring of each guanine is forced to move further away from the cis guanine and closer to the cis amine (‘6-out’ relationship). If the cis amine has an N-H on the same side of the platinum coordination plane as the O6 of the guanine, an H-bond could possibly be formed.101 Between HT conformers, the DHT1 L appears to be a major component in the case of adducts with the (S,R,R,S)-bip ligand, which in theory can provide two N-Hs for H-bond formation with the O6 of the two guanines (this can be seen in Figure 5.7, in which, however, the two guanines are not tethered).106 The LHT2 R conformer can also form two guanine O6/NH cis amine H-bonds in adducts of (R,S,S,R)bip (this can also be seen in Figure 5.7, although here the two guanines are not tethered); however, at acidic pH the LHT2 R conformer is a minor form compared to the HH1 R and HH2 R (the latter only for the d(GpG) adduct) conformers, thus indicating that the LHT2 R conformer is intrinsically less stable. The LHT2 R conformer concentration increases at high pH, and this could be attributed to O6 becoming a better hydrogen-bond acceptor when N1 is deprotonated.103 5.6.2 Multiple Conformers in Cisplatin Adducts with Tethered Guanine Bases The cis-(NH3)2Pt(GpG) complex is probably very dynamic, undergoing rapid rotation about the Pt-N7 bonds. The H8 signals of cis-(NH3)2Pt(GpG) are relatively sharp (∼10 and 4 Hz for 5′-G H8 and 3′-G H8, respectively) at 21 °C but broader (∼20 and 5 Hz, respectively) at 5 °C.103 Therefore, exchange between conformer classes or between variants within a conformer class must be moderately fast at 21 °C and slower at 5 °C. Comparison to spectral data for bip-Pt(GpG) indicates that cis-(NH3)2Pt(GpG) exists as a mixture of variants from both the DHT1 and HH1 conformer classes;103 thus, the most likely explanation for the broadening is exchange between conformer classes. Analysis of the NMR shifts and CD signal

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shapes indicates that at low pH cis-(NH3)2Pt(GpG) exists mostly as the HH1 R conformer, as suggested previously in the literature;123 however, other conformers account for perhaps one-third of the complex. At high pH, the conformer distribution changes. Although it appears to be reasonable that the HH1 L variant is now a more abundant form, it is not the exclusive form. Other forms likely to be present include the DHT1 form.103 As found for the GpG adduct, cis-(NH3)2Pt(d(GpG)) also exists as a dynamic mixture of various forms of comparable populations, in contrast to the adducts with longer oligonucleotides, which exist mainly as the HH1 L variant.106 In the solid state, cis-(NH3)2Pt(d(pGpG)) crystallizes with an equal population of HH1 R and HH1 L variants.132 We believe that a mixture of these two variants also exists in solution. In addition, the computed anti/syn DHT1 model of cis-(NH3)2Pt(d(GpG)) is also structurally very similar to that computed for the analogous bip adduct. Furthermore, we computed that this conformer has one of the lowest energies of the cis-(NH3)2Pt(d(GpG)) models and we have found this form in many adducts containing a wide variety of carrier ligands; therefore, it is most likely that the DHT1 form is also part of the dynamic mixture of cis-(NH3)2Pt(d(GpG)) conformers. 5.6.3 Wrapping of the Positively Charged Metallic Core by Flanking Residues in Cisplatin Adducts with Single-Stranded Oligonucleotides With the exception of very simple species such as cis-(NH3)2Pt(d(GpG)), an unusual but characteristic feature of larger oligonucleotides crosslinked at N7 by platinum(II) is a large difference (∼1 ppm) in the H8 signals, with the 3′-G H8 signal downfield in single-stranded adducts. The guanine orientation dictated by the sugar–phosphate backbone is responsible for the shift difference.108 The same remarkable difference in H8 chemical shifts of the HH conformer was found in Me2dab and bip-PtG2 complexes with untethered guanines, suggesting that interligand interactions in the coordination sphere may dictate guanine orientations comparable to those induced by the sugar–phosphate backbone of the GpG moiety within larger oligonucleotides. On the basis of the unusual 31P NMR signals observed in the LPt– d(TCTCGGTCTC) adducts (L = cis-(NH3)2 or en), it was concluded that this G/G crosslinked single strand is a distorted coil with the 5′ residues ‘wrapped around’ the Pt binding site.133 The 5′-region distortion is probably caused both by electrostatic interactions between the negatively charged nucleotide sequences flanking the crosslink and the cationic [cis-(NH3)2Pt]2+ or [en-Pt]2+ moieties and by hydrogenbond formation between the phosphate and the Pt-NH groups; the resulting local distortion at phosphodiester groups explains the rather unusual 31P NMR shifts.134 Such ‘wrapping’ of the cationic [cis-(NH3)2Pt]2+ moiety by flanking residues results in more stable coils and in local ‘melted’ regions of longer DNA, thereby decreasing the Tm of DNA.134,135 Treatment of the self-complementary dodecamer d(A1T2G3G4G5T6A7C8C9C10 A11T12) (G3) under conditions in which G3 is in the duplex (DS) form with cisplatin or en-PtCl2 gives single-stranded (SS) products: G3/G4 adducts in a coil form and G4/G5 adducts in a hairpin form stabilized by a favourable A7-G4 stacking interaction at the loop–stem junction (Figure 5.14).67,136–138

Retro Models Applied to Adducts 161

Figure 5.14 Schematic representation of the reaction of G3 (d(A1T2G3G4G5T6A7C8C9C10A11 T12)) with: (A) platinum complexes with secondary nitrogen donors having steric bulk (small loops close to N) or (B) platinum compounds without steric bulk on the nitrogen atoms (Ref. 139 for related work)

The ‘wrapped’ SS coil form becomes less stable in bip-Pt–G3 and Me2dab-Pt– G3 adducts because the hydrophobic and bulky bip and Me2dab carrier ligands prevent the flanking regions of the negative DNA strand from approaching the cationic Pt centre as closely as these regions can approach the Pt in adducts with less hindered bis-ammine or primary diamine ligands. These effects involving the bip and Me2dab ligands while destabilizing the ‘wrapped’ form, favour the duplex form; therefore, the bip-Pt–G3 and Me2dab-Pt-G3 adducts with the crosslink in a HH1 conformation favour the duplex form (Figure 5.14).139 This is particularly true for (R,S,S,R) configuration of the diamine which induces the characteristic R canting of platinated guanines in duplexes. In the case of (S,R,R,S) configuration of the diamine, the duplex appears to be in equilibrium with a small amount of a single-stranded (most likely coil) form. The facts that the (S,R,R,S) chirality favours L canting and that this canting is characteristic of coils suggest a reasonable explanation for this result. An important outcome of this investigation is that the conformation and annealing propensity of SS oligonucleotides with G/G intrastrand crosslinks can be modulated by the stereochemistry of the platinum carrier ligands. 5.6.4 Flexibility in Cisplatin Adducts with Double-Stranded Oligonucleotides A few X-ray structures of double-stranded (DS) oligonucleotides containing a cisA2Pt moiety (A2 = two ammines or a diamine) crosslinking two G residues of the same strand72,73,140,141 or of opposite strands74 have been reported. In the intrastrand

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view from the major groove

view from the minor groove

platinum

nitrogen oxygen

carbonium phosphorus

Figure 5.15 (Plate 9) Central portion (four base-pairs) of oligonucleotides containing G/G intrastrand (top) and interstrand (bottom) crosslinks (based on data from Refs 114 and 143, respectively) (See colour plate section)

crosslinks (intra-CL) the cis-A2Pt moiety is located in the major groove, and the six-membered rings of the two guanines are on the same side of the platinum coordination plane (head-to-head, HH, conformation, Figure 5.15). In contrast, when platinum crosslinked G residues of opposite strands (inter-CL), the cis-A2Pt moiety

Conclusions and Perspectives

163

is located in the minor groove, and the six-membered rings of the two guanines are on opposite sides of the platinum coordination plane (head-to-tail, HT, conformation, Figure 5.15). Solution NMR data have been interpreted by analogy with the results of the X-ray investigations.46,70,114,142,143 A detailed description of these adducts appears in some excellent reviews.144–146 However, even in the case of DS oligonucleotides, which are, per se, somewhat rigidly structured, some dispute exists as to the fine structure of more dynamic regions, and the contributions of the sugar–phosphate backbone and of interligand interactions in determining the overall stereochemistry remain rather uncertain. For instance, no two NMR studies on duplexes have led to the same proposed structure46,68,70,107,147 and the diverse proposed solution structures generally lack the features found in the X-ray structure of a 16-mer bound to rat HMG1.148 These variations in structure may be caused by spectral complications arising from dynamic motion centred at the crosslink. For instance, the dihedral angle between crosslinked guanines has been found to span all values between 25 ° and 75 °. However, in a relatively recent study of a 9-mer duplex, an NMR-based structural model with many of the elements of the X-ray structure of a 16-mer bound to rat HMG1 was reported,114 and this 9-mer has many spectral features common to duplexes with a pyrimidine–G–G–pyrimidine sequence. If, as appears to be the case, the structure of crosslinked double-stranded oligonucleotides is far from being rigid, it is likely that the conformation is deeply influenced by ‘weak’ interligand interactions, which are difficult to detect, but which can result in a different biological activity. This appears to be the case for oxaliplatin, [Pt(oxalato)(dach)] (dach = 1,2-diaminocyclohexane), for which only the R,R enantiomer ((R,R)-dach carrier ligand) has been approved for clinical use.141,144 We also investigated the mutagenic activity of several platinum compounds with chiral diamines and found that the S,S enantiomer was always a far greater mutagenic agent than the R,R form, particularly in the case of dach and dab adducts (dab = 2,3diaminobutane).149 In an insightful analysis the distortion of double-stranded oligonucleotides modified at a single site by dach-Pt or dab-Pt residues of different chiralities (either R,R or S,S) was monitored by chemical probes. It was found that different diamine enantiomers induced different types of distortion and, as a consequence of these structural differences, the affinity of the platinated oligonucleotides for proteins (e.g. HMG proteins) was different, as well as their processing by repair systems.150,151

5.7 Conclusions and Perspectives In general, adducts of cisplatin (and cisplatin analogues) with nucleotides and SS and DS oligonucleotides are highly dynamic systems that are difficult to investigate with conventional techniques. In most cases intermolecular interactions will affect the solid-state conformation; solution data are generally interpreted on the basis of a single model even when the system is likely to be a mixture of rapidly interconverting forms. Retro models have proven to be particularly useful in unravelling the

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existence of different conformations, and in the most favourable cases have also allowed the full characterization of their stereochemistry. Moreover, several interligand interactions, some of which had remained unrecognized for a long time, have been revealed and a hierarchy among them has been established. Our analysis, based on solution results with adducts containing specially designed carrier ligands, has allowed us to rationalize computational, solution-state, and solid-state results on cis-(NH3)2Pt crosslink adducts in a more consistent manner than previous interpretations based on the cis-(NH3)2Pt adducts themselves. This success verifies the value of the retro modelling approach. In addition, by using retro models we found dramatic differences in the relative stability of different forms of oligonucleotides (coil, hairpin and duplex) dependent on the carrier ligand. We believe that in the future the use of retro models for the stabilization of single conformers and the characterization of their stereochemistry will play an important role in the interpretation of experimental data pertaining to dynamic systems and in the validation of results coming from theoretical investigations.

Acknowledgements G. Natile gratefully acknowledges the collaboration with Prof. Dr L. G. Marzilli (Louisiana State University) which, over a period of nearly two decades, has led to the major discoveries presented in this article. Thanks are also due to the many students and postdoctoral researchers who have contributed and whose names are listed in the references as coauthors of previous papers. Support and sponsorship from European COST Actions D39/0004/06 ‘Structure, recognition, and processing of DNA damage by antitumour metal-based compounds’, the Italian Ministry for University and Research (MUR, Rome), the University of Bari and the Interuniversity Consortium for Research in the Chemistry of Metal Ions in Biological Systems (C.I.R.C.M.S.B.) are also acknowledged.

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124. Pasini, A.; De Giacomo, L.; Aspects of the stereochemistry of bis-guanosine-5′monophosphate diamine platinum complexes; Inorg. Chim. Acta, 1996, 248, 225–230. 125. Song, B.; Oswald, G.; Bastian, M.; Sigel, H.; Lippert, B.; Extent of the acidification by N7-coordinated cis-diammine-platinum(II) on the acidic sites of guanine derivatives; Metal-Based Drugs, 1996, 3, 131–141. 126. Carlone, M.; Fanizzi, F.P.; Intini, F.P.; Margiotta, N.; Marzilli, L.G.; G. Natile; Influence of carrier ligand NH hydrogen bonding to the O6 and phosphate group of guanine nucleotides in platinum complexes with a single guanine ligand; Inorg. Chem., 2000, 39, 634–641. 127. Carlone, M.; Marzilli, L.G.; Natile, G.; Platinum complexes with NH groups on the carrier ligand and with only one guanine or hypoxanthine derivative. Informative models for assessing relative nucleobase and nucleotide hydrogen-bond interactions with amine ligands in solution; Inorg. Chem., 2004, 43, 584–592. 128. Benedetti, M.; Marzilli, L.G.; Natile, G.; Rotamer stability in cis- Pt(diA)G2 complexes (diA = diamine derivative and G = guanine derivative) mediated by carrier-ligand amine stereochemistry as revealed by circular dichroism spectroscopy; Chem. Eur. J., 2005, 11, 5302–5310. 129. Ranaldo, R.; Margiotta, N.; Intini, F.P.; Pacifico, C.; Natile, G.; Conformer distribution in (cis-1-4-DACH)bis(guanosine-5′-phosphate)platinum(II) adducts: a reliable model for DNA adducts of antitumoural cisplatin; Inorg. Chem., 2008, 47, 2820–2830. 130. Chottard, J.C.; Girault, J.P.; Chottard, G.; Lallemand, J.Y.; Mansuy, D.; Interaction of cisdiaquodiammineplatinum dinitrate with ribose dinucleoside monophosphates; J. Am. Chem. Soc., 1980, 102, 5565–5572. 131. Lebwohl, D.; Canetta, R.; Clinical development of platinum complexes in cancer therapy: an historical perspective and an update; Eur. J. Cancer, 1998, 34, 1522–1534. 132. Sherman, S.E.; Gibson, D.; Wang, A.H.J.; Lippard, S.J.; Crystal and molecular structure of cis-Pt(NH3)2 d(pGpG), the principal adduct formed by cis-diamminedichloroplatinu m(II) with DNA; J. Am. Chem. Soc., 1988, 110, 7368–7381. 133. Kline, T.P.; Marzilli, L.G.; Live, D.; Zon, G.; Investigations of platinum amine induced distortions in single- and double-stranded oligodeoxyribonucleotides; J. Am. Chem. Soc., 1989, 111, 7057–7068. 134. Pilch, D.S.; Dunham, S.U.; Jamieson, E.R.; Lippard, S.J.; Breslauer, K.J.; DNA sequence context modulates the impact of a cisplatin 1,2-d(GpG) intrastrand crosslink on the conformational and thermodynamic properties of duplex DNA; J. Mol. Biol., 2000, 296, 803–812. 135. Poklar, N.; Pilch, D.S.; Lippard, S.J.; Redding, E.A.; Dunham, S.U.; Breslauer, K.J.; Influence of cisplatin intrastrand crosslinking on the conformation, thermal stability and energetics of a 20-mer DNA duplex; Proc. Natl. Acad. Sci. USA, 1996, 93, 7606–7611. 136. Kline, T.P.; Marzilli, L.G.; Live, D.; Zon, G.; NMR studies of an oligonucleotide with an unusual structure induced by platinum anticancer drugs; Biochem. Pharmacol., 1990, 40, 97–113. 137. Villanueva, J.M.; Jia, X.; Yohannes, P.G.; Doetsch, P.W.; Marzilli, L.G.; Cisplatin (cis2+ Pt(NH3)2Cl2) and cis-Pt ( NH 3 )2 ( H 2O)2 intrastrand crosslinking reactions at the telomere GGGT DNA sequence embedded in a duplex, a hairpin, and a bulged duplex: Use of Mg2+ and Zn2+ to convert a hairpin to a bulged duplex; Inorg. Chem., 1999, 38, 6069–6080. 138. Yohannes, P.G.; Zon, G.; Doetsch, P.W.; Marzilli, L.G.; DNA hairpin formation in adducts with platinum anticancer drugs: gel electrophoresis provides new information and a caveat; J. Am. Chem. Soc., 1993, 115, 5105–5110. 139. Beljanski, V.; Villanueva, J.M.; Doetsch, P.W.; Natile, G.; Marzilli, L.G.; Marked dependence on carrier-ligand bulk but not on carrier-ligand chirality of the duplex versus single-strand forms of a DNA oligonucleotide with a series of G-Pt(II)-G intrastrand crosslinks modeling cisplatin-DNA adducts; J. Am. Chem. Soc., 2005, 127, 15833–15842.

References

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140. Spingler, B.; Whittington, D.A.; Lippard, S.J.; 1,2-d(GpG) intrastrand crosslink formed by oxaliplatin with duplex DNA: a crystallographic study; J. Inorg. Biochem., 2001, 86, 440–440. 141. Spingler, B.; Whittington, D.A.; Lippard, S.J.; 2.4 Å crystal structure of an oxaliplatin 1,2-d(GpG) intrastrand crosslink in a DNA dodecamer duplex; Inorg. Chem., 2001, 40, 5596–5602. 142. Huang, H.; Zhu, L.; Reid, B.R.; Drobny, G.P.; Hopkins, P.B.; Solution structure of a cisplatin-induced DNA interstrand crosslink; Science, 1995, 270, 1842–1845. 143. Paquet, F.; Perez, C.; Leng, M.; Lancelot, G.; Malinge, J.-M.; NMR solution structure of a DNA decamer containing an interstrand crosslink of the antitumour drug cis-diammi nedichloroplatinum(II); J. Biomol. Struct. Dyn., 1996, 14, 67–77. 144. Hambley, T.W.; The influence of structure on the activity and toxicity of Pt anti-cancer drugs; Coord. Chem. Rev., 1997, 166, 181–223. 145. Reedijk, J.; Improved understanding in platinium antitumour chemistry; J. Chem. Soc. Chem. Commun., 1996, 801–806. 146. Jamieson E. R. Lippard, S J.; Structure, recognition and processing of cisplatin-DNA adducts; Chem. Rev., 1999, 99, 2467–2498. 147. Herman, F.; Kozelka, J.; Stoven, V.; Guittet, E.; Girault, J.P.; Huynh-Dinh, T.; Igolen, J.; Lallemand, J.Y.; Chottard, J.C.; A d(GpG)-platinated decanucleotide duplex is kinked. An extended NMR and molecular mechanics study; Eur. J. Biochem., 1990, 194, 119–133. 148. Ohndorf, U.M.; Rould, M.A.; He, Q.; Pabo, C.O.; Lippard, S.J.; Basis for recognition of cisplatin-modified DNA by high-mobility-group proteins; Nature, 1999, 399, 708–712. 149. Coluccia, M.; Correale, M.; Giordano, D.; Mariggiò, M.A.; Moscelli, S.; Fanizzi, F.P.; Natile, G.; Maresca, L.; Mutagenic activity of some platinum complexes with monodentate and bidentate amines; Inorg. Chim. Acta, 1986, 123, 225–229. 150. Malina, J.; Kasparkova, J.; Natile, G.; Brabec, V.; Recognition of major DNA adducts of enantiomeric cisplatin analogs by HMG box proteins and nucleotide excision repair of these adducts; Chem. Biol., 2002, 9, 629–638. 151. Kasparkova, J.; Vojtiskova, M.; Natile, G.; Brabec, V.; Unique properties of DNA interstrand crosslinks of antitumour oxaliplatin and the effect of chirality of the carrier ligand; Chem. Eur. J., 2008, 14, 1330–1341.

6 Role of DNA Repair in Antitumour Effects of Platinum Drugs Viktor Brabec and Jana Kašpárková

6.1 Introduction Understanding the molecular and biochemical mechanisms associated with the biological effects of the existing platinum agents provides important insights for designing new and more efficient platinum-based drugs. There is a large body of experimental evidence suggesting that the success of platinum complexes in killing tumour cells results from their ability to damage DNA by forming various types of covalent adducts.1–4 Hence the research into DNA interactions with platinum antitumour drugs has predominated.5,6 However, cells have developed efficient tools to correct any chemical changes in DNA so that the genetic information is retained uncorrupted. Agents that damage DNA involve: (i) ionizing and ultraviolet radiation; (ii) highly reactive oxygen radicals produced during normal cellular respiration, as well as by other biochemical pathways; (iii) chemicals in the environment, such as various hydrocarbons, including some found in cigarette smoke, some plant, fungal and microbial products, e.g. the aflatoxins produced in mouldy peanuts and not least (iv) chemicals used in chemotherapy, especially in chemotherapy of cancer (see Table 6.1). Major types of DNA damage are: (i) all four of the bases in DNA (A, T, C, G) which can be covalently modified at various positions. One of the most frequent is the loss of an amino group (‘deamination’) resulting, for example, in a C being Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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Table 6.1 Examples of chemotherapeutic agents that damage DNA Drug Bleomycin Busulfan Chlorambucil Cyclophosphamide Melphalan Mitomycin Carboplain Cisplatin Oxaliplatin Dactinomycin (actinomycin D) Daunorubicin (daunomycin) Doxorubicin Epirubicin Idarubicin Irinotecan Mitoxantrone Nogalamycin Proflavin Berenil Hoechst 33258 Lexitropsins (netropsin, distamycin and analogues)

Mechanism of action Cuts DNA strands between GC or GT Alkylating agent forming inter- and/or intrastrand crosslinks

Form intrastrand and intersand cross-links Inhibit DNA and RNA synthesis by intercalating between base pairs of the DNA/RNA strand, thus preventing the replication of rapidly growing cancer cells; they also create iron-mediated oxygen free radicals that damage the DNA and cell membranes; are also capable of inhibiting topoisomerase II enzymes, preventing DNA from relieving torsional stress during replication Inhibit DNA and RNA synthesis by binding noncovalently to the minor groove of DNA

converted to a U; (ii) mismatches of the normal bases due to a failure of proofreading during DNA replication; (iii) breaks in the backbone which can involve both strands (double-strand break) or be limited to only one strand (single-strand break) (ionizing radiation is a frequent cause, but some chemicals produce breaks as well); (iv) crosslinks – covalent linkages can be formed between bases on the same DNA strand (‘intrastrand’) or on the opposite strands (‘interstrand’), between two DNA double helices or between DNA and proteins. Several chemotherapeutic drugs used against cancers including antitumour platinum drugs crosslink DNA. Damage to the DNA molecules is identified by the cell, which uses a collection of mechanisms to correct this damage. This process, called DNA repair, is defined as a biological process during which alterations in the chemistry of DNA (DNA damage) are removed and the integrity of the genome restored. The broader field is now more appropriately referred to as the field of biological responses to DNA damage, and in this context even the term ‘DNA damage’ has been broadened to include phenomena such as the arrest of DNA synthesis in the absence of defined DNA damage.7 The adducts formed by platinum antitumour drugs cause structural damage to DNA and can alter or eliminate the cell’s ability to replicate and transcribe the gene that the affected DNA encodes. These lesions also induce potentially harmful mutations in the cell’s genome, which affect the survival of its daughter cells after it undergoes mitosis. After DNA is modified by platinum drugs, cellular repair systems recognize this damage. DNA repair is not perfect, however.This means that over

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time the repair process will fail and a cell will enter one of the following states: (i) apoptosis or cellular suicide and necrosis; these are processes by which the cell destroys itself; (ii) senescence, a process whereby the cell becomes irreversibly dormant; (iii) uncontrolled cellular division; this can result in tumour formation and cancer. Thus, DNA repair has a significant role in modulating cytotoxicity of platinum drugs. The interruption of pathways that are involved in repairing platinated DNA represents a powerful approach to cancer treatment. It is therefore not surprising that repair of damage to DNA induced by platinum antitumour drugs and its downstream effects in tumour cells is, at present, a very active area of molecular pharmacology. The purpose of this review is to highlight recent discoveries related to repair of DNA damage induced by anticancer platinum drugs. The topics discussed do not involve mechanisms underlying repair of DNA damage by platinum compounds in bacterial or yeast systems.

6.2 Human DNA Repair Systems All macromolecules in a living cell, i.e., nucleic acids, proteins, lipids and carbohydrates can be damaged when the cells are exposed to deleterious agents, including antitumour metal-based drugs. A major defence against damage to cells is DNA repair. Although other biomacromolecules may be damaged and subject to repair, the significance and exceptionality of DNA repair in cellular processes is accentuated by the fact that only failure to repair DNA damage has fatal consequences for the cells. If other cellular molecules are damaged, they can be degraded and newly synthesized via transcription and translation. Thus, for instance, damage to cellular proteins is usually not corrected since, due to the large number of different chemical modifications, it seems to be chemically and evolutionally impossible to develop the required set of enzymes to correct damage to proteins. Therefore, very often protein degradation is the only way to remove such damaged proteins from the protein pool. Ribozyme mediated strategies exist that can be used to amend damaged genetic instructions at the RNA level.8–10 An obvious limitation of RNA repair is, however, the fact that repair of RNA only results in transient production of corrected genetic instructions, as the damaged DNA template is not amended. DNA repair, which is present in all organisms examined, including bacteria, yeast, drosophila, fish, amphibians, rodents and humans, is involved in processes that minimize cell death, mutations, replication errors, persistence of DNA damage and genomic instability. All organisms must keep their DNA intact and free of lesions to obtain faithful transmission of genetic information. There are several different DNA repair pathways in mammalian cells, as described in the sections 6.2.1–6.2.4. 6.2.1 Direct Reversal of Damage This type of DNA repair is relatively rare and involves chemical reversal of damage. It does not involve breakage of the phosphodiester backbone and is specific to the

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type of damage. Only a few types of DNA damage are repaired in this way, particularly pyrimidine dimers resulting from exposure to ultraviolet (UV) light and alkylated guanine residues that have been modified by the addition of methyl or ethyl groups at the O6 position of the purine ring. Enzymes that catalyse direct repair reactions are widespread in both prokaryotes and eukaryotes, including humans. This type of DNA repair has not been observed for DNA damaged by antitumour metal-based drugs. 6.2.2 Excision Repair Although direct repair is an efficient way of dealing with particular types of DNA damage, excision repair is a more general means of repairing a wide variety of chemical alterations to DNA, including those induced by platinum antitumour drugs. In excision repair, the damaged DNA is recognized and removed, either as free bases or as nucleotides. The resulting gap is then filled in by synthesis of a new DNA strand, using the undamaged complementary strand as a template. Three types of excision repair – base-excision repair (BER), nucleotide-excision repair (NER) and mismatch repair (MMR) – enable cells to cope with a variety of different kinds of DNA damage. Base Excision Repair Common small lesions, especially methylated or oxidized bases, or uracil in DNA resulting from spontaneous deamination of cytosine are corrected by this type of DNA repair. An altered base is specifically recognized by one of many specialized glycosylases. The recognition process may be sensitive to enhanced DNA flexibility or to decreased base-pair stability as well as to steric effects like protrusions in the grooves. The glycosylase removes the offending base, giving an abasic site or AP site (apurinic/apyrimidinic site). The common feature of the glycosylases is that they tend to flip out the base and do chemistry on the extrahelical base. Excision of a damaged base and subsequent processing of the resulting single-strand break lead to an intermediate, containing a 3′-hydroxyl, suitable for priming strand resynthesis by a polymerase (Table 6.2). The majority of BER proceeds through the so-called ‘short-patch’ subpathway in which a single nucleotide is removed and replaced. In this pathway (Figure 6.1), DNA resynthesis is carried out by polymerase b (Table 6.2), a member of the X family of polymerases (this DNA polymerase has the unique ability to repair single-stranded DNA gaps smaller than six nucleotides; it fills single nucleotide gaps in DNA produced by the BER pathway of mammalian cells).11 DNA ligase I or III completes the repair by sealing DNA ends.12 Nucleotide Excision Repair In NER a small region of the strand surrounding the damage is removed from DNA as an oligonucleotide. The small gap left in the DNA helix is filled by the sequential action of DNA polymerase and DNA ligase. NER is generally thought of as a repair system for bulky adducts, that recognizes a wide range of substrates, such as damage caused by UV irradiation and chemical agents, including platinum anticancer drugs.

Human DNA Repair Systems 179 Table 6.2 The survey of currently known human DNA polymerases and their function.a89,154 Polymerase name Polymerase a (synonymes are DNA primase, RNA polymerase) Polymerase b Polymerase g Polymerase d Polymerase e Polymerases h, i, k, and Rev1 Polymerase z Polymerase q, l, j, s, and m Others, but their nomenclature is ambiguous

Polymerase proposed function Acts as a primase (synthesizing a RNA primer), and then as a DNA polymerase elongating that primer with DNA nucleotides Is implicated in repairing DNA (BER; single-strand break repair) Replicates mitochondrial DNA Is the main polymerase on the lagging strand in eukaryotes, it is highly processive and has 3′ → 5′ exonuclease activity Is the primary leading strand DNA polymerase in eukaryotes, and is also highly processive and has 3′ → 5′ exonuclease activity Y-family DNA polymerases, TLS B-family DNA polymerase, TLS Not well characterized

a None of the eukaryotic polymerases can remove primers (5′ → 3′ exonuclease activity); that function is carried out by other enzymes. Only the polymerases that deal with elongation (g, d and e) have proofreading ability (3′ → 5′ exonuclease).

NER is a complex process (Figure 6.2) in which basically the following steps can be distinguished: (i) recognition of a DNA lesion; (ii) separation of the double helix at the DNA lesion site; (iii) single strand incision at both sides of the lesion; (iv) excision of the lesion-containing single stranded DNA fragment; (v) DNA repair synthesis to replace the gap and (vi) ligation of the remaining single stranded nick. In step (i) two modes of recognition can be distinguished (Figure 6.2): repair of lesions over the entire genome, referred to as global genome NER (GG-NER), and repair of transcription-blocking lesions present in transcribed DNA strands, hence called transcription-coupled NER (TC-NER). GG-NER removes damage in both transcribed and untranscribed DNA strands in active and inactive genes throughout the genome. This pathway employs several damage-recognition proteins that scan the genome and recognize distortions in double-helical DNA. Unlike GGNER, TC-NER is initiated, not by specific recognition of the DNA damage site by proteins in the NER pathway, but rather by stalling RNA polymerase II. Human GG-NER has been characterized at the biochemical level in considerable detail.13 Six repair factors, XPA, RPA, XPC-RAD23B, TFIIH, XPG and XPFERCC1, are necessary and sufficient to remove damage from DNA. A current model for human NER is as follows: XPA, RPA and XPC-RAD23B locate the damage site; whether the primary damage recognition factor is RPA (or XPA/RPA complex) or XPC-RAD23B is unclear, inconsistent findings in regard to the order of the recognition assembly have been reported. Another NER protein thought to cooperate in DNA damage recognition is XPE protein14 although several studies

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Figure 6.1 Base excision repair mechanism in mammalian cells

have raised some doubts about the role of XPE in excision repair.14–16 The recognition assembly recruits the TFIIH transcription/repair factor, which contains six polypeptides including helicases XPB and XPD that unwind the DNA around the damage site; the XPG and XPF-ERCC1 subunits are responsible for the 3′ and 5′ dual incisions, respectively. The dual-incision leads to the removal of a singlestranded fragment of DNA with a single-strand gap of 25∼30 nucleotides. The resulting gap in DNA is filled by DNA polymerase d or e (Table 6.2) by copying the undamaged strand. Proliferating cell nuclear antigen (PCNA) assists the DNA polymerase in the reaction, and replication protein A (RPA) protects the other DNA strand from degradation during NER. Finally, DNA ligase I seals the nicks to finish NER.17,18 Mismatch Repair MMR provides several genetic stabilization functions; it corrects errors of DNA replication, ensures the fidelity of genetic recombination and participates in the earliest steps of checkpoint and apoptotic responses to several classes of DNA damage. Hence, a primary function of the MMR pathway is to correct persistent DNA replication errors and avoid the accumulation of deleterious mutations. MMR

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Figure 6.2 Nucleotide excision repair mechanism in mammalian cells

repairs base–base mismatches and insertion–deletion loops generated during DNA replication and recombination.19 The MMR system is also of biomedical interest because MMR-deficient tumour cells are resistant to certain cytotoxic chemotherapeutic drugs, including cisplatin.20,21 MMR has evolved to correct errors of DNA polymerases that escape their 3′ → 5′ exonuclease proofreading activity. These errors, mismatches, can be identified in DNA because they fail to form Watson–Crick base pairs. However, because

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neither nucleotide is damaged or modified, it is not obvious which strand carries the correct genetic information and which carries the error; thus, mismatch repair cannot be accomplished by a mechanism such as BER or NER, which simply excise the damaged base, or a short DNA fragment containing the damage, respectively (see above). Unlike BER and NER, postreplicative MMR has to be targeted exclusively to the newly synthesized strand, which carries, by definition, the erroneous genetic information.21 However, as the parent and daughter strands cannot be distinguished at the site of the mispair, this has to happen elsewhere. The most important features of the current models of the molecular mechanism of MMR are the following:22 The MMR process (Figure 6.3) is initiated by the binding of the hMutSα (hMSH2/ hMSH6 heterodimer), which undergoes an ATP-driven conformational change and recruits the hMLHl/hPMS2 heterodimer (also in an ATP-dependent manner). This ternary complex can translocate in either direction along the DNA. Importantly, the binding of a mismatch or modified DNA base(s) by the hMSH2/hMSH6 heterodimer, or by the hMSH2/hMSH6/hMLHl/hPMS2 complex, does not bring about DNA incision at this site, nor does it lead to damage excision. When it encounters a strand discontinuity that may be bound by a proliferating cell nuclear antigen, loading of an exonuclease initiates degradation of the nicked strand towards the mismatch. Notably, the exonucleolytic degradation of DNA takes place only if a pre-existing strand discontinuity is present in the vicinity. If the exonuclease dissociates before it reaches the mismatch, the single-stranded gap is stabilized by RPA. Loading of the second hMutSα/hMLHl/hPMS2 complex at the mismatch stimulates a second round of exonucleolytic degradation. This process is repeated until the mismatch is removed. Also importantly, the modified DNA bases are not removed during the DNA degradation step unless they are present in the strand containing the discontinuity. The RPA-stabilized single-stranded gap is filled in by the replicative polymerase and the remaining nick can be sealed by DNA ligase.23,24 6.2.3 Repair of Double-Strand Breaks Double-strand breaks, in which the phosphodiester backbones of both strands in the double helix are interrupted, are particularly hazardous to the cell because they can lead to genome rearrangements. Two mechanisms exist to repair double-strand breaks: nonhomologous end joining (NHEJ) and homologous recombination repair (HHR) (also known as template-assisted repair). Nonhomologous End Joining DNA NHEJ is a predominant pathway of DNA double-strand break repair in mammalian cells, and defects in the pathway cause radiosensitivity at the cellular and whole-organism levels. NHEJ is referred to as ‘nonhomologous’ because the break ends are directly ligated without the need for a homologous template, in contrast to homologous recombination (see below), which requires a homologous sequence to guide repair. NHEJ typically utilizes the short homologous DNA sequences in the single-stranded overhangs that are often present at the ends of double-strand breaks and are used to promote restorative repair. When these overhangs are com-

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Figure 6.3 Mismatch repair mechanism in mammalian cells

patible, NHEJ almost always repairs the break accurately, with no sequence loss.25 This mechanism does not have quality control on the basis of sequence homology and, hence, is not error-free, and can lead to deletions and inversions. Imprecise repair leading to loss of nucleotides can also occur, but is much less common. Interestingly, cisplatin inhibits repair of double-strand breaks by NHEJ.26–28

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Figure 6.4 Nonhomologous end-joining mechanism in mammalian cells

The major NHEJ pathway (Figure 6.4) relies on a set of core proteins. The two DNA ends of the double-strand break are recognized and bound by the ring-shaped heterodimer Ku70/Ku80,which recruits the DNA-dependent protein kinase catalytic subunit (DNA-PKcs).29 The assembled DNA-dependent protein kinase (DNAPK) holoenzyme then exhibits serine-threonine protein kinase and DNA end-bridging activities.30 Among other functions, the kinase activity regulates DNA end access to processing enzymes like the DNA-PKcs-associated Artemis nuclease.31,32 Finally, the XRCC4/DNA ligase IV complex is responsible for the ligation step.33 Another core NHEJ factor is Cernunnos-XLF, a factor with a predicted structural similarity to XRCC4 that has been identified as an XRCC4-interacting protein.34 DNA polymerases l and m (Table 6.2) fill gaps during NHEJ.35 Homologous Recombination Repair HRR acts on double-strand breaks generated at broken replication forks or occurring within replicated DNA. Recombinational repair requires the presence of an identical or nearly identical sequence to be used as a template for repair of the break. The enzymatic machinery responsible for this repair process is nearly identical to the machinery responsible for chromosomal crossover during meiosis. This pathway allows a damaged chromosome to be repaired using a sister chromatid or a homologous chromosome as a template.

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The HRR process (Figure 6.5) requires the assembly of multienzymatic complexes.36 These complexes include the Rad51 family members such as Rad51, Rad54, Rad51B, Rad51C, Rad51D, Xrcc2 and Xrcc3.37 To initiate the repair of doublestrand breaks in DNA, RAD51 needs to bind to the single-stranded DNA that is produced by nucleolytic resection at the break site. The protein shows a weak specificity for binding single-stranded DNA compared with double-stranded DNA, but this specificity is enhanced by interactions with another key recombination protein, which is known as RAD52. Invasion of a resected end of the double-strand break into duplex DNA takes place in the RAD51 filament and requires the binding of a high energy nucleotide cofactor such as ATP.

6.2.4 DNA Damage Bypass Even when DNA repair and cell cycle checkpoint control are fully functional, some DNA lesions often persist through replication of the genome. Factors that contribute to the persistence of DNA damage include: (i) high levels of damage; (ii) poorly repaired lesions; (iii) inefficiently repaired genomic regions and (iv) DNA damage incurred during the S phase of the cell cycle. Since many lesions that persist despite DNA repair and cell cycle checkpoints hamper or counteract the replication apparatus, cells have evolved a damage tolerance system to allow complete replication in the presence of DNA damage. There are two reasons why it is important for the cell to be able to move replication forks past unrepaired damage. First, long-term blockage of replication forks leads to cell death. Second, replication of damaged DNA provides a sister chromatid that can be used as template for subsequent repair by homologous recombination. DNA damage bypass or translesion synthesis (TLS) is a DNA damage tolerance process that allows the DNA replication machinery to replicate past DNA lesions (Figure 6.6). It involves exchanging regular DNA polymerases by specialized translesion polymerases (Table 6.2), often with larger active sites that can facilitate the insertion of bases opposite damaged nucleotides. The polymerase switching is thought to be mediated by, among other factors, the post-translational modification of the replication processivity factor – PCNA. Many of the TLS enzymes (Table 6.2) belong to the recently described ‘Y-family’ of DNA polymerases. By possessing a spacious preformed active site, these enzymes can physically accommodate a variety of DNA lesions and facilitate their bypass. Flexible DNA-binding domains and a variable binding pocket for the replicating base pair further allow these TLS polymerases to select specific lesions to bypass and favour distinct nonWatson–Crick base pairs. Consequently, TLS polymerases tend to exhibit much lower fidelity (higher propensity to insert wrong bases) than the cell’s regular polymerases when copying normal DNA, which results in a dramatic increase in mutagenesis. Occasionally this can be beneficial, but it often speeds the onset of cancer in humans. However, many TLS polymerases (Table 6.2) are extremely efficient at inserting correct bases opposite specific types of damage. For example, polymerase h mediates error-free bypass of lesions induced by UV irradiation, whereas polymerase z introduces mutations at these sites. From a cellular perspective, risking the introduction of point mutations during TLS may be preferable to resorting to more drastic

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Figure 6.5 Homologous recombination repair mechanism in mammalian cells

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Figure 6.6 Mechanism of translesion synthesis in mammalian cells

mechanisms of DNA repair, which may cause gross chromosomal aberrations or cell death. Cells use both transcriptional and posttranslational regulation to keep these low-fidelity polymerases under strict control and limit their access to a replication fork.38 Lesion bypass directly utilizes the damaged template. Conceptually, lesion bypass can be divided into two steps: (i) nucleotide incorporation opposite the lesion (i.e., translesion synthesis), followed by (ii) extension of DNA synthesis. After a short stretch of extension, normal DNA synthesis by the replication apparatus can then resume. Significantly, lesion bypass can be either error-free, whereby the correct

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nucleotide opposite the damage is predominantly incorporated, or error-prone, whereby an incorrect nucleotide is frequently incorporated opposite the damage. Consequently, error-free lesion bypass is a mutation-avoiding mechanism, and errorprone lesion bypass is a mutation-generating mechanism. Several proteins, including novel Y-family polymerases that have been recently identified in a multitude of organisms, are involved in DNA damage bypass pathways. Humans have four such (Y-family) polymerases (Table 6.2) – polymerase k, polymerase i, polymerase h, and Rev1 – each with a unique DNA damage bypass and fidelity profile. Polymerase h, for example, is unique in its ability to replicate through UV-induced cyclobutane pyrimidine dimers, while polymerase k is inefficient at replicating through a T-T dimer, but can readily extend from mispaired termini. Polymerase i is perhaps the most unusual with varied efficiencies and fidelities opposite different template bases. Notably, polymerases involved in TLS make up an appreciable fraction of all known eukaryotic DNA polymerases (Table 6.2). Some of the TLS polymerases are capable of replicating accurately past certain types of DNA damage. Rev1 does not have a Greek name since it’s not considered a true DNA polymerase; it can insert only one nucleotide at a time, usually in a template-independent fashion. Polymerases i and k are related to polymerase h but have somewhat different bypass specificities and are not usually error-free. Polymerase q is especially competent at bypassing AP sites, where it inserts an A residue. Since most AP sites are consequences of purine loss, and A is a purine base, repair by polymerase q is frequently error-free.

6.3 Specific Binding of Repair Proteins to DNA Modified by Antitumour Platinum Compounds A number of proteins that are components of various cellular DNA repair systems bind to DNA modified by antitumour platinum drugs specifically (Table 6.3). When these proteins are absent, repair of DNA damage by platinum complexes is diminished; sensitivity of cells to the platinum drug is enhanced, which is consistent with the hypothosis that these proteins modulate biological effects of these metallodrugs. Some of these proteins initiate specific cell signalling pathways. Recognition of DNA adducts of antitumour platinum compounds by DNA repair systems has mostly been examined in the case of DNA adducts of cisplatin and its analogs cis-diamminecyclobutanedicarboxylatoplatinum(II)] (carboplatin) and [(1R,2R-diamminocyclohexane)oxalatoplatinum(II)] (oxaliplatin). Cisplatin binds to DNA, forming mainly intrastrand crosslinks between adjacent purine residues (∼90%) (1,2-GG or 1,2-AG intrastrand crosslinks).39,40 Other minor adducts are 1,3-GXG intrastrand crosslinks (X = A, C, T), interstrand crosslinks (∼6% in linear DNA) preferentially between guanine residues in the 5′GC/5′-GC sequence41 and monofunctional lesions. The adducts formed by cisplatin in DNA affect its secondary structure.42 The formation of major 1,2-crosslinks of cisplatin locally unwinds DNA duplex and bends it towards the major groove, expos-

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Table 6.3 Examples of human proteins involved in DNA repair that were shown to bind specifically to DNA modified by antitumour platinum compounds Protein 3-methyladenine DNA glycosylase (AAG) XPA RPA XPC-HR23B RNA polymerase II PARP-1 hMutSα (heterodimer of hMSH2 and hMSH6) Ku80 SSRP1

Proposed function

Reference

BER NER NER, HRR NER TC-NER TC-NER MMR NHEJ Transcription elongation

97 51 51,53 60 64–67 69 70,71 76,78 95

ing a wide, shallow minor groove surface to which several classes of proteins bind. The interstrand crosslink also induces several irregularities in DNA.43 The crosslinked guanine residues are not paired with hydrogen bonds to the complementary cytosines, which are located outside the duplex. The cis-diammineplatinum(II) bridge resides in the minor groove and the double helix is locally reversed to a lefthanded, Z-DNA-like form, markedly locally unwound and bent towards the minor groove. The 1,3-GXG intrastrand crosslink formed by cisplatin locally unwinds and bends the helix axis towards the major groove, but in contrast to 1,2-intrastrand adducts, DNA is locally denatured and flexible at the site of the 1,3-adduct.44 The monofunctional adducts of cisplatin distort DNA in a sequence-dependent manner.45,46 These distortions disturb stacking interactions in double-helical DNA, slightly unwind the duplex, but no intrinsic bending is induced by these adducts. Direct analogues of cisplatin, such as carboplatin and oxaliplatin, produce adducts similar to those produced by the parent drug, though different in their relative rates of formation.47–49 6.3.1 Base Excision Repair System The human 3-methyladenine DNA glycosylase (AAG) is a BER enzyme that removes a number of damaged bases from DNA, including adducts formed by some chemotherapeutic agents. AAG readily recognizes cisplatin adducts, such as 1,2-GG, 1,2-AG and 1,3-GTG intrastrand crosslinks. 1,N-6-ethenoadenine, which is repaired efficiently by AAG, is recognized considerably more weakly than cisplatin adducts.96 6.3.2 Nucleotide Excision Repair System The NER proteins that exhibit enhanced affinity to cisplatin-modified DNA are mainly those involved in the first step in repair pathways, i.e. in damage recognition. The repair proteins that probably have attracted the most attention are those that are absent in patients suffering from the NER deficiency characteristic of the disease

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Xeroderma pigmentosum (XP). The minimal factors necessary for removal of damaged nucleotides also include XPA and RPA which are among the major damage-recognition proteins involved in the early stages of NER. XPA (32 kDa) and RPA (70, 34 and 14 kDa subunits) are able to bind damaged DNA independently, although RPA interaction stimulates XPA binding to damaged DNA. XPA belongs among proteins that bind preferentially to bent or kinked DNA duplexes.50 Interestingly, XPA protein binds with considerable selectivity to the rigid double-stranded kink induced by the 1,2-GG intrastrand crosslink of cisplatin.51 In contrast, XPA fails to recognize the site of strong duplex destabilization generated by the same crosslink, but formed by the bifunctional dinuclear platinum complex [{trans-PtCl(NH3)2}2H2N(CH2)4NH2]Cl251 that fails to impose a rigid kink, but, instead, increases DNA flexibility in a nondirectional manner.52 RPA, on the other hand, recognizes the helical destabilization caused by the dinuclear complex more effectively than the cisplatin-induced kink. RPA binds with a relatively low affinity to double-stranded DNA containing the 1,2-GG intrastrand crosslink.53 The preference of RPA for the more helix destabilizing crosslink54 is consistent with previous studies suggesting that RPA is recruited to damaged double-stranded DNA through interactions with single-stranded sites.53,55 It has been suggested51 that XPA in conjunction with RPA constitutes a regulatory factor that monitors DNA bending and unwinding. In addition, it has been demonstrated56 that the particular manner in which XPA and RPA proteins align themselves with respect to the 1,3-GNG intrastrand crosslink involves their binding in close proximity to the adduct, with XPA in contact with both the platinated and unplatinated strands of DNA, and RPA binding preferentially to the unplatinated strand. It has also been shown57 that a direct interaction between RPA and XPA proteins facilitates the assembly of a preincision complex during the NER processing of DNA containing a 1,2-GG intrastrand crosslink. A ternary complex of RPA and XPA both bound to DNA containing the 1,2-GG intrastrand crosslink displays modest specificity for platinated versus unplatinated DNA. In addition, the RPA–XPA complex exhibits a greater affinity for binding cisplatin-damaged duplex as compared to the RPA or XPA proteins alone. It has also been demonstrated57 that the role of XPA is to stabilize the doublestranded DNA structure by inhibiting the strand separation activity of RPA. XPA is also believed to participate in DNA damage recognition of cisplatin adducts, whereas it has no affinity for DNA adducts of clinically ineffective transplatin.58 The effects of the lesions induced by single, site-specific 1,2-GG or 1,3-GTG intrastrand adducts of cisplatin on the energetics of DNA have been examined as well.59 The results have confirmed that DNA bending is the specific determinant responsible for high-affinity interactions of XPA with damaged DNA. An additional important factor is a change of thermodynamic stability of DNA induced by the damage. These results confirm that RPA preferentially binds to DNA distorted in such a way that hydrogen bonds between complementary bases are interrupted. RPA also binds to nondenatured distortions in double-helical DNA, but the affinity of RPA to these distortions is insensitive to alterations of thermodynamic stability of damaged DNA. XPC-HR23B displays an enhanced binding affinity for 1,3-GTG intrastrand crosslinks compared to unmodified DNA.60 In addition, XPC-hHR23B associates

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with the 1,2-GG and 1,3-GXG intrastrand crosslinks with similar rates. The more rapid rate of dissociation of XPC-HR23B from the 1,2-GG compared to the 1,3GXG intrastrand crosslink is likely a result of the lack of localized melting of the duplex DNA containing the 1,2-GG intrastrand adduct. RPA is one of the proteins that belongs to the initial damage-sensing factors initiating NER (see above). The relative binding affinities of RPA to the different cisplatin adducts correlate with the repair of the adducts observed in an in vitro NER assay;61 RPA has higher affinity for the 1,3-GTG intrastrand than for the 1,2GG intrastrand crosslink of cisplatin, and the 1,3-GTG adduct is more efficiently repaired by the human exonuclease than the 1,2-GG adduct.53 Thus, binding of RPA to damaged DNA is a sensitive indicator to predict whether mammalian NER could be effective in the removal of damaged nucleotides. BBR3464 is the novel phase II antitumour polynuclear platinum drug ([(trans-PtCl(NH3)2)2(m-trans-Pt-(NH3)2(NH2 (CH2)6NH2)2)]4+), which forms on DNA so-called long-range intra- and interstrand crosslinks. The effect of 1,4-GG intrastrand and interstrand crosslinks of BBR3464 on RPA binding was examined62 (the platinated bases in 1,4-crosslinks are separated by two intervening base pairs). Interestingly, RPA did not bind to DNA containing interstrand crosslinks of BBR3464. Thus, these results predict very low efficiency of the mammalian NER pathway to remove the interstrand crosslinks of BBR3464, subsequently proved by in vitro experiments.62,63 TC-NER is initiated by stalling RNA polymerase II. Therefore, cisplatin DNA adduct processing by RNA polymerase II has been investigated.64–67 It was found that RNA polymerase II stalls in front of a cisplatin intrastrand crosslink because it does not pass a translocation barrier that impairs delivery of the bulky crosslink into the active site. In vitro assays indicate that a considerable fraction of stalled RNA polymerase II proteins remains strongly associated with DNA damaged by cisplatin following RNA polymerase II arrest.66 RNA polymerase II stalled in front of a cisplatin adduct (1,3-GTG intrastrand crosslink) serves as a decoy to sequentially recruit TFIIH, XPA, RPA, XPG and XPF repair factors in an ATP-dependent manner. This RNA polymerase II/repair complex allows the ATP-dependent removal of the cisplatin lesion.65 In conclusion, the anticancer drug cisplatin forms DNA intrastrand crosslinks that stall RNA polymerase II, and in this way cisplatin triggers TC-NER.67 In addition, recent observations demonstrate a role for poly-(ADPribose)polymerase-1 (PARP-1) in TC-NER68 and PARP-1 binding to 1,2-GG intrastrand crosslinks in nuclear cell extracts.69 Thus, the latter observations indicate that poly(ADP-ribosylation) activity may be crucial for cellular NER of platinated DNA.

6.3.3 Mismatch Repair System Similarly to the case of NER, the enhanced affinity to DNA damaged by cisplatin is exhibited mainly by the MMR proteins involved in damage recognition, i.e. by one of two heterodimers of MutS homologues, hMutSa (heterodimer of hMSH2 and hMSH6) or hMutLa (hMLH1 and hPMS2 heterodimer). The purified hMSH2 protein binds to DNA containing a single 1,2-GG intrastrand crosslink.70 Moreover, hMSH2

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displays selective affinity for DNA modified by cisplatin, but not for DNA modified by transplatin70 and preferentially recognizes cisplatin-modified DNA over oxaliplatinmodified DNA.71 The hMutSα heterodimer recognizes the 1,2-GG intrastrand crosslink of the cisplatin adduct, but it has no affinity for the 1,3-GTG intrastrand crosslink of transplatin.72 The binding affinity of hMutSα for the 1,2-GG intrastrand crosslink of cisplatin is enhanced in duplexes containing a thymine incorporated opposite the 5′ or the 3′ guanine of the cross-link.73,74 The data suggest that hMutSα may interact with cellular platinum-DNA lesions (especially with intrastrand crosslinks superimposed on a mismatch73) and influence the DNA repair and signalling pathway, although a detailed mechanism of these processes is not known. 6.3.4 Binding of Proteins Involved in the Nonhomologous End-Joining System DNA-PK is a core protein of the NHEJ pathway and binds to DNA modified by cisplatin. Its K80 subunit, which is responsible for the binding,75 also has a high affinity to the major adduct of cisplatin, the 1,2-GG intrastrand crosslink, which is only twofold lower than its affinity to DNA ends.76 DNA adducts of cisplatin do not alter the ability of Ku to bind DNA ends, but do impair the translocation of Ku proteins along DNA,77 resulting in reduced affinity of DNA-PKc to the Ku-DNA complex, and reduced kinase activity.78 6.3.5 Homologous Recombination Repair System The Rad51-guided HRR system plays an important role in the recognition and repair of DNA interstrand crosslinks. It is not unexpected that cells deficient in the HRR pathway become hypersensitive to cisplatin as well as to other interstrand crosslinking agents. The repair of interstrand crosslinks has been shown to involve components of the NER, HRR and TLS pathways.79 However, the manner by which HRR proteins specifically bind platinum adducts is unclear. 6.3.6 Processing of DNA Modified by Antitumour Platinum Compounds by Translesion Synthesis DNA Polymerases Several TLS DNA polymerases (Table 6.2) bypass 1,2-GG intrastrand crosslinks of cisplatin.80–85 DNA polymerases that bypass cisplatin adducts in vitro include DNA polymerases b, m and h, whereas polymerases a, i, k and l are unable to perform TLS past platinum adducts.80,82,83,86 Each DNA polymerase displays a distinct specificity in its lesion-bypass properties, including bypass ability, fidelity and extension ability. For example, DNA polymerase h bypasses platinum adducts most efficiently in error-free TLS,81,85 whereas polymerase m is the most error-prone enzyme. Moreover, two DNA polymerases often work together to complete TLS.87 Although polymerase z is unable to bypass certain DNA lesions, including those by platinum agents, the enzyme has the ability to extend TLS once nucleotides are inserted opposite DNA adducts by other polymerases.88,89 Little is known about the TLS past platinum interstrand crosslinks, although a single DNA polymerase is not likely to

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be able to bypass this lesion. It has been suggested, however, that TLS may occur during the repair of interstrand crosslinks.79,90 Interestingly, polymerase h bypasses 1,2-GG crosslinks of oxaliplatin more readily than 1,2-GG crosslinks of cisplatin.91 This suggests that the conformations of cisplatin- and oxaliplatin-GG adducts are different and that these conformational differences have an influence on the ability of distantly related DNA polymerases to perform translesion synthesis. 6.3.7 Binding of Other DNA Repair Proteins The high-mobility-group (HMG) SSRP1 protein (structure-specific recognition protein 1) is a member of a conserved chromatin-remodelling complex (FACT/ DUF/CP) implicated in DNA replication, basal and regulated transcription, and also in DNA repair.92,93 SSRP1 is the first HMG-domain protein found to bind selectively to DNA adducts of cisplatin.94,95 The role of this protein in repair of DNA adducts of platinum compounds is, however, unknown.

6.4 Repair of DNA Damage by Antitumour Platinum Compounds 6.4.1 Base excision Repair The presence of cisplatin adducts in the reactions inhibits the excision of 1,N-6ethenoadenine by BER DNA glycosylase AAG. AAG readily recognizes cisplatin intrastrand adducts while the 1,N-6-ethenoadenine adduct, which is repaired efficiently by AAG, is recognized considerably more weakly. Despite the affinity of AAG for cisplatin adducts, AAG is unable to remove any of these adducts from DNA. It has been suggested that cisplatin adducts could titrate AAG away from its natural substrates, resulting in higher mutagenesis and/or cell death because of the persistence of AAG substrates in DNA.96 Another intriguing possibility is that AAG binds to cisplatin adducts and, because it cannot remove them, it conveys the adducts to the NER pathway through the interaction of AAG with the hHR23 proteins.97 Thus the interaction between AAG and DNA adducts of cisplatin can increase the efficiency of DNA repair and, in the case of overexpression of this protein, could lead to cisplatin resistance.97,98 Hence, cisplatin may exhibit a synergistic effect in potentiating the toxic effects of agents that are substrates for AAG. 6.4.2 Nucleoide Excision Repair A major pathway used by human cells for the removal of platinum adducts from DNA is NER. Typical experiments demonstrating NER of DNA adducts of platinum complexes were performed using linear DNA fragments of ∼150 base pairs containing a platinum adduct in the centre of one strand. To allow for determination of excision activity in human or rodent cell extracts, the modified sequences con-

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tained a 32P-labeled phosphate at the phosphodiester bond near the lesion on its 5′ side. Following incubation with cell extract or a reconstituted system containing highly purified NER factors, damage-containing oligonucleotides are released as radiolabelled DNA fragments that are visualized after gel electrophoresis and autoradiography. Incubation in mammalian cell extracts or a reconstituted system containing highly purified NER factors demonstrated that both major 1,2- and minor 1,3-intrastrand crosslinks of cisplatin are efficiently repaired, the 1,3-crosslink being repaired much more efficiently.61 Both crosslinks bend and unwind DNA to a similar extent, but in contrast to 1,2-intrastrand crosslinks, the 1,3-intrastrand adduct of cisplatin locally denatures and induces a flexibility at the site of the 1,3-adduct.44 Hence, the differential NER can be attributed to the different DNA structures induced by the 1,2- and 1,3-intrastrand cisplatin adducts. Denatured base pairs and locally enhanced flexibility induced in DNA by platinum complexes may be factors enhancing recognition of platinum adducts by NER systems. On the other hand, in the extracts from human HeLa cells, the 1,2-AG intrastrand crosslink is excised more efficiently than the 1,2-GG intrastrand adduct61 although these 1,2-intrastrand crosslinks distort the DNA in a similar way (DNA unwinding and bending are similar6,99,100 and hydrogen bonds in the distorted base pairs are preserved). NER of 1,2-, but not of 1,3-intrastrand crosslinks, is blocked upon addition of an HMG-domain protein (HMG = high mobility group).101 Several mechanisms have been considered for how HMG-domain proteins might modulate the sensitivity of cells to cisplatin.61 One prominent hypothesis is that HMG-domain proteins block cisplatin–DNA adducts from the damage recognition needed for repair. Hence, in the cells where this shielding mechanism is responsible for their sensitivity to cisplatin, 1,2- and not 1,3-intrastrand crosslinks of this metallodrug are most likely candidates for genotoxic lesions relevant to the antitumour effects of cisplatin. An in vitro NER of a site-specific cisplatin interstrand crosslink has also been studied using mammalian cell-free extracts containing HMG-domain proteins at levels insufficient to block NER of the 1,2-intrastrand adducts.61 Repair of the interstrand crosslink formed by cisplatin was not detected. Similarly, in cell strains derived from patients with Fanconi’s anaemia, NER of cisplatin-interstrand crosslinks is not observed, although NER can readily occur in these cells.102,103 Fanconi’s anaemia cells have been described as being extremely sensitive to interstrand crosslinking agents, and it was suggested that this high sensitivity to cisplatin could be explained by the incapability of these cells to repair cisplatin interstrand crosslinks.104 On the other hand, repair of these lesions has been detected with the aid of a repair synthesis assay, which measures the amount of new DNA synthesized after damage removal in whole-cell extracts.105 In this way, however, the repair could also result from a mechanism different from that of NER. DNA interstrand crosslinks pose a special challenge to repair enzymes because they involve both strands of DNA and therefore cannot be repaired using the information in the complementary strand for resynthesis. Quite recently the processing of stalled forks caused by DNA interstrand crosslinks has been proposed to be an important step in initiating mammalian interstrand crosslink repair. It has been demonstrated106 that the XPFERCC1 complex makes an incision 5′ to a psoralen interstrand crosslink on Y-shaped DNA and that the XPF-ERCC1 complex generates an interstrand crosslink-specific incision on the 3′-side of this lesion. The interstrand crosslink-specific 3′ incision,

Repair of DNA Damage by Antitumour Platinum Compounds 195

along with the 5′ incision, on the crosslinked Y-shaped DNA results in the separation of the two crosslinked strands (the unhooking of the interstrand crosslink) and the induction of a double-strand break near the crosslinked site. These results implicate the XPF-ERCC1 complex in initiating interstrand crosslink repair by unhooking the interstrand crosslink, which simultaneously induces a double-strand break at a stalled fork. That this mechanism asserts itself also in repair of interstrand crosslinks of cisplatin and other platinum complexes awaits further experimental confirmation. The differential recognition of the 1,2-GG intrastrand crosslink of cisplatin and dinuclear platinum complex [{trans-PtCl(NH3)2}2H2N(CH2)4NH2]Cl2 with divergent effects on DNA conformation (see above) by XPA and RPA prompted testing of NER activity in response to these two platinum lesions.51 Incubation of linear DNA fragments containing the platinum crosslink in HeLa cell extract demonstrated that 1,2-GG intrastrand crosslinks of both platinum complexes were able to elicit oligonucleotide excision. These results are consistent with the view, and support the hypothesis that XPA-RPA constitutes a universal sensor of nucleotide damage that operates through indirect readout of DNA conformations. XPA-RPA is guided by the synergism between two different DNA binding domains: XPA recognizes abnormal rigid bending of the deoxyribose-phosphate backbone, whereas RPA recognizes base pair disruption. These two nucleic acid interaction modules are used to monitor integrity of the Watson–Crick helix by a double-check probing mechanism that confers an extremely wide recognition capacity for structural distortions. Cisplatin also forms DNA–protein cross-links107 The DNA–protein cross-links formed by cisplatin inhibit removal of these ternary lesions from DNA by NER systems more effectively than plain DNA intrastrand or monofunctional adducts. Thus, the bulky DNA–protein crosslinks formed by cisplatin represent a more distinct and persisting structural motif recognized by the components of downstream cellular systems processing DNA damage considerably different from that of the plain DNA adducts of this metallodrug. In addition, in cell-free media, cisplatin forms DNA–protein crosslinks considerably more effectively than the clinically ineffective transplatin. This implies that there is a positive correlation between the efficiency of mononuclear bifunctional platinum complexes to form DNA–protein crosslinks and their antitumour effects. Thus, these results are consistent with previous findings demonstrating that DNA–protein crosslinks persist longer in cells exposed to cisplatin than in those exposed to the chemotherapeutically inactive trans analogue,108–110 which suggests relevance of such lesions to the antitumour effects of cisplatin. The failure to remove bulky DNA–protein crosslinks has been suggested to be due to the steric hindrance caused by the size of the crosslinked protein,which may interfere sterically with the assembly of the mammalian NER system. Thus, the results demonstrating failure of the mammalian NER system to remove the DNA–Pt–protein crosslinks107 reinforce the view that DNA–protein crosslinks formed by platinum drugs may be among the critical lesions relevant to their antitumour effects. In eukaryotic cells, DNA is packed and the smallest structural unit of eukaryotic DNA packaging, is the nucleosome. The nucleosome is made up of about 146 base pairs of DNA and four pairs of proteins called histones. Histones H2A, H2B, H3 and H4 form the core of the nucleosome, around which the DNA is wrapped,

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while histone H1 sits on the base of the nucleosome at the junction between nucleosome DNA and linker DNA, extending along the DNA into the linker region.111 Nucleosomes modulate many cellular processes so that it is of interest to know how NER of DNA adducts of platinum compounds is affected when these adducts are present in nucleosomes. Nucleosomes were prepared containing either a site-specific 1,2-GG or 1,3 GTG intrastrand crosslink of cisplatin.112 It was found that the nucleosome significantly inhibited NER, the inhibition being more pronounced in the case of the 1,3intrastrand crosslink. Interestingly, excision from native nucleosomal DNA was higher than the level observed with recombinant material. This result reveals that posttranslational modification of histones can modulate NER from damaged chromatin. Despite its success, cisplatin has several disadvantages: it is active against a limited range of human tumours, can cause severe side effects and its administration may result in acquired resistance. The drawbacks, coupled with the toxicity of cisplatin and its analogues, have been the impetus for the development of platinum drugs with improved pharmacological properties and a broader range of antitumour activity. Antitumour activity of platinum drugs is multifactorial in nature and also includes contributions from differential DNA repair mechanisms. Therefore it is of interest to know whether NER and other repair pathways discriminate between cisplatin and novel platinum drugs that exhibit altered biological effects. A study of this kind was focused on NER of DNA adducts formed by a third generation antitumour platinum drugs oxaliplatin [(1R,2R-diamminocyclohexane) oxalato-platinum(II)] and satraplatin [bis-acetatoamminedichlorocyclohexylamine platinum(IV), JM216].113 Interestingly, the types of DNA lesions generated by the three platinum drugs, cisplatin, oxaliplatin and satraplatin, are repaired in vitro with similar kinetics by the mammalian NER pathway. This result is consistent with reports that major DNA adducts of cisplatin, oxaliplatin and satraplatin cause similar distortions in DNA.114,115 Besides oxaliplatin ([Pt(R,R-DACH)]2+, DACH = 1,2-diaminocyclohexane), another enantiomeric form of this complex exists, [Pt(S,S-DACH)]2+. Whereas conformational alterations induced in DNA by the major adduct of [Pt(R,R-DACH)]2+ are not markedly different from those induced by the adduct of cisplatin, conformational alterations induced by the major adduct formed by [Pt(S,S-DACH)]2+ in the TGGT sequence are different.115 The major differences resulting from the modification by the two enantiomers consist in sequence-dependent thermodynamic destabilization and conformational distortions. Interestingly, the 1,2-GG intrastrand crosslinks of [Pt(R,R-DACH)]2+ bind to HMG-domain proteins with a similar affinity as the same crosslink of cisplatin.115 In contrast, the crosslink formed by [Pt(S,S-DACH)]2+ in the TGGT sequence binds to the HMG-domain protein with a noticeably lower affinity. Similar results have also been obtained with DNA modified by [Pt(DAB)]2+ (DAB = 2,3-diaminobutane) enantiomers (which are closely related to [Pt(DACH)]2+ enantiomers).116–118 Intrastrand crosslinks between neighbouring guanine residues of both enantiomers of [Pt(DAB)]2+ were efficiently removed from DNA by NER in an in vitro

Repair of DNA Damage by Antitumour Platinum Compounds 197

assay using mammalian or rodent cell-free extracts.117 Consistent with the different affinity of the intrastrand crosslinks of both enantiomers to HMG-domain proteins, addition of HMG-domain protein only blocked NER of the crosslinks of [Pt(R,RDAB)]2+, whereas NER of the crosslinks formed by [Pt(S,S-DAB)]2+ in the TGGT sequence remained unaffected. Thus, these results suggest that very fine structural modifications, such as those promoted by enantiomeric ligands in bifunctional platinum(II) compounds, can modulate the ‘downstream effects’, such as specific protein recognition by DNA processing enzymes and other cellular components. More specifically, as a consequence of enantiomorphism of carrier amine ligands of cisplatin analogues, their major DNA crosslinks not only exhibit different conformational features, but are also processed differently by cellular components, including NER. It may be also suggested that these differences are associated with different antitumour and other biological activities of the two enantiomers. The original empirical structure–activity relationships considered the trans isomer of cisplatin and other transplatin analogues to be inactive.119 However, several new analogues of transplatin, which exhibit a different spectrum of cytostatic activity including activity in tumour cells resistant to cisplatin have been identified. Examples are analogues containing heterocyclic amine ligands, aliphatic ligands or iminoether groups.3,120,121 The molecular mechanism of antitumour activity of transplatinum compounds has been extensively investigated. Unlike cisplatin or transplatin, modification of DNA by antitumour trans compounds leads to monofunctional and bifunctional intra- and interstrand crosslinks in different proportions. In addition, structure of intrastrand and interstrand crosslinks of antitumour trans complexes and conformational distortions induced by these adducts in DNA are mostly fundamentally different from that of the crosslinks of cisplatin and parent transplatin. Thus, it is not surprising that cellular processing of DNA adducts of antitumour transplatin analogues, including NER, is also different. The interstrand crosslinks of these trans-platinum compounds are not removed from DNA in the same way as the corresponding adducts of cisplatin. In contrast to cisplatin, intrastrand crosslinks of antitumour trans-platinum analogues are removed from DNA by NER with markedly lower efficiency than the major adducts of cisplatin.122,123 Hence, a general property of DNA adducts of these new antitumour trans-platinum compounds is that they can persist for a considerably long time in cells, which is a prerequisite for the toxicity of these lesions in tumors sensitive to these transplatin analogs, without being protected from the recognition and attack of the NER components. An intriguing example of how NER modulates the antitumour effects of transplatinum compounds is the case of trans-[PtCl2(E-iminoether)2]. The replacement of both NH3 groups in transplatin by an iminoether ligand results in a marked enhancement of its cytotoxicity. It is more cytotoxic than its cis congener and exhibits significant antitumour activity, including activity in cisplatin-resistant tumour cells.124 The new trans compound (trans-[PtCl2(E-iminoether)2]) forms mainly stable monofunctional adducts at guanine residues when reacted with DNA.125 These adducts locally distort DNA, are not recognized by HMG-domain proteins and are readily removed from DNA by the NER system. The monofunctional adducts of trans-[PtCl2(E-iminoether)2] readily crosslink proteins, which markedly enhances

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the efficiency of this adduct to terminate DNA polymerization in vitro and to inhibit removal of this adduct from DNA by the NER system.126 Hence, DNA–protein ternary crosslinks produced by trans-[PtCl2(E-iminoether)2] persist considerably longer than the noncrosslinked monofunctional adducts, which potentiates toxicity toward tumour cells. Thus, trans-[PtCl2(E-iminoether)2] belongs to a class of platinum antitumour drugs in which activation of the trans geometry is associated with an increased efficiency to form DNA–protein ternary crosslinks. Polynuclear platinum compounds comprise a unique class of anticancer platinum agents with distinct chemical and biological properties different from mononuclear platinum drugs.127 The lead compound is BBR3464, a trinuclear, bifunctional DNA binding agent. Quite surprisingly, in comparison with cisplatin, enhanced cellular uptake of the charged polynuclear platinum compounds has been demonstrated,128,129 but the enhanced uptake is not sufficient to explain the increased toxicity of BBR3464 in tumour cells.130 The interactions of antitumour polynuclear platinum compounds with target DNA (for review see, for instance, ref. 127) are distinct from the mononuclear-based cisplatin family and, indeed, unlike those of any DNA-damaging agent in clinical use. The ability of BBR3464 to induce long-range delocalized intraand interstrand crosslinks,131 which are not produced by conventional mononuclear platinum compounds, suggests that BBR3464 may escape, at least in part, the classical mechanism of cisplatin resistance related to DNA damage recognition and repair. The crosslinks of BBR3464 distort the DNA conformation. In contrast to distortions induced by the major crosslinks of cisplatin, the crosslinks of BBR3464 do not extensively unwind and rigidly bend DNA. Hence they are not substrates for damaged-DNA binding proteins, such as HMG-domain proteins.62,63 On the other hand, while intrastrand adducts of BBR3464 are readily removed from DNA by the NER systems, the interstrand crosslinks are not. It has been suggested62,63 that interstrand, and not intrastrand, crosslinks of BBR3464 could persist for a sufficiently long time in cells to potentiate toxicity of this drug.

6.4.3 Mismatch Repair Recent observations support the view that another cellular repair mechanism, such as MMR, can affect the antitumour efficiency of cisplatin,132,133 and that dysfunction of this type of DNA repair may result in the resistance of tumour cells to cisplatin, or in drug tolerance.134 The function of the MMR is to scan newly synthesized DNA and remove mismatches that result from nucleotide incorporation errors made by the DNA polymerases. To explain cisplatin tolerance, it is assumed that replication bypass of DNA adducts of cisplatin leads to mutations. During MMR, the strand to be corrected is nicked, a short fragment containing the mismatch is excised, and a new DNA fragment is synthesized. The MMR system always replaces the incorrect sequence in the daughter strand, which would leave the cisplatin adduct unrepaired. This activity initiates a futile cycle. During DNA synthesis to replace the excised short fragment, the DNA polymerases would again incorporate mutations followed by attempts to remove them. The repeated nicks in DNA formed at each ineffective cycle of repair

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could trigger a cell death response. Thus, MMR recognition of cisplatin adducts on DNA may trigger a programmed cell death pathway rendering MMR-proficient cells more sensitive to DNA modification by cisplatin than MMR-deficient cells. In other words, loss of MMR would then increase the cell’s ability to tolerate lesions formed on DNA by cisplatin. The MMR is likely to act mainly in the postreplicative phase, when the highest concentration of mispairs is expected in the newly synthesized DNA. This would be in contrast to NER, which probably acts independently of the cell cycle. Thus, rapidly proliferating tumour cells would be more prone to futile repair attempts capable of triggering cell death. Although the level of resistance to cisplatin that accompanies loss of MMR is relatively small, loss of MMR has been shown to be sufficient to account for the failure of treatment with cisplatin in several model systems.135–137 Nevertheless, a direct connection between the MMR pathway and cytotoxicity of cisplatin has not been established. 6.4.4 Nonhomologous End-Joining Radiation exposure can produce many types of DNA damage, including doublestrand breaks. In a local multiply damaged site, where an adduct of cisplatin occurs close to a single-strand break, the damage may be initially repaired by NER. However, during the dual incision step, a double-strand break, which is the most lethal type of a cell damage, is produced. Alternatively, if an adduct of cisplatin is close to a double-strand break, it could physically block cisplatin removal and repair by the NER pathway, or the cisplatin adduct could hinder the double-strand break repair pathways by restricting the access of the protein complexes involved. Another possibility is that the NER and proteins involved in repair of double-strand breaks may compete for access to the local multiply damaged site. Investigations into the efficiency of NER have shown that NER is less efficient if cisplatin adducts are near to a double-strand break due to binding of the heterodimer Ku70/80 at the DNA ends.138 In addition, a decreased amount of NER proteins binds to the lesion if a double-strand break is present.139,140 Double-strand break NHEJ, which requires DNA-PK, is also inhibited by cisplatin-damaged DNA in cell extracts.26,27 This process may in part explain the radiosensitizing effect of cisplatin administered during concurrent chemoradiation. 6.4.5 Homologous Recombination Repair Several studies have shown that HRR plays a role in the repair of DNA damage by cisplatin in mammalian cells.21,141,142 HRR of unrepaired cisplatin adducts at stalled replication forks appears to be a mechanism of cellular resistance to cisplatin in NER-deficient and MMR-deficient cells. In addition to its function in NER and its ability to uncouple cisplatin interstrand crosslinks, the ERCC1/XPF structurespecific nuclease has a role in the repair of cisplatin adducts related to recombination processing.143 Homologous recombination might be initiated prior to excision of interstrand crosslinks of cisplatin. Because of its probable role in HRR of DNA adducts of cisplatin, ERCC1 is an attractive therapeutic target, especially in MMRdeficient cancers, where increased recombinational bypass of cisplatin lesions

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enhances resistance to this drug. It has been demonstrated144 that downregulation of ERCC1 significantly sensitizes the MMR-deficient prostate cancer cell line to cisplatin, where inhibition of the NER pathway had no effect. In addition, a recent study21 suggests that homologous recombination status might represent a novel prognostic marker and possibly also a therapeutic target.

6.5 Implications for Design of Antitumour Platinum Compounds The occurrence of inherent and acquired resistance to platinum is a major problem that undermines efforts to effectively treat cancer. This limitation in cancer treatment can be overcome by elucidating the mechanisms underlying resistance to platinum drugs and then developing ways to treat resistant tumours effectively or prevent its occurrence.145 DNA repair is a major resistance mechanism to platinumbased chemotherapy. Evidence for increased repair of platinum–DNA damage in resistant cancer cells has been demonstrated by a variety of cellular assays. In human cells, major intrastrand crosslinks of cisplatin are removed from DNA mainly by the NER system.61 Importantly, the repair of major intrastrand crosslinks of cisplatin may be blocked by high-mobility-group (HMG) domain proteins, and perhaps by other proteins that bind to damaged DNA,3,97,146,147 causing drug resistance to be lower. Thus, the ability of cancer cells to recognize damage induced in DNA by platinum antitumour drugs and initiate DNA repair is an important mechanism for therapeutic resistance and has a negative impact upon therapeutic efficacy. Pharmacological inhibition of DNA repair, therefore, has the potential to enhance efficiency of antitumour chemotherapy. It implies that design of new antitumour platinum drugs should focus on those new platinum complexes that form major adducts with DNA that are difficult to remove by DNA repair systems. This strategy has been employed by designing and synthesizing antitumour dinuclear platinum complexes [(cis-{Pt-(NH3)2})2(m-OH)(m-pz)]2+ (pz = pyrazolate)148 or dinuclear azole-bridged platinum compounds.149–152 These new platinum complexes exhibit higher toxic effects in some tumour cell lines and form 1,2-GG instrastrand crosslinks, which bend DNA markedly less than cisplatin. It has been suggested that these minor structural changes are not recognized by the DNA repair proteins and in this way escape repair. This assumption must, however, await further experimental confirmation. It cannot be excluded that the bulkiness of the adduct in addition to its propensity to induce only very weak conformational distortions is an important factor involved in the initiation of DNA repair. Moreover, the use of inhibitors of DNA repair or DNA damage signalling pathways also appears to provide an opportunity to enhance or alter the antitumour effects of platinum agents and to target the genetic differences that exist between normal and tumour tissue as well.153 On the other hand, use of DNA repair inhibitors or their combination with such anticancer therapies is also likely to enhance the risk of mutagenic DNA damage and hence the risk of secondary cancer. Thus, concepts for antitumour strategies based on DNA repair inhibitors must await further studies.

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Acknowledgements This work was supported by the Ministry of Education of the Czech Republic (MSMT LC06030, 6198959216, ME08017, OC08003), the Academy of Sciences of the Czech Republic (Grants 1QS500040581, KAN200200651, AV0Z50040507 and AV0Z50040702), the Grant Agency of the Academy or Sciences of the Czech Republic (IAA400040803) and the Grant Agency of the Ministry of Health of the Czech Republic (NR8562-4/2005). J.K. is the international research scholar of the Howard Hughes Medical Institute. The authors also acknowledge that their participation in the EU COST Action D39 enabled them to exchange regularly the most recent ideas in the field of anticancer metallodrugs with several European colleagues.

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96. Kartalou, M.; Samson, L.D.; Essigmann, J.M.; Cisplatin adducts inhibit 1,N-6-ethenoadenine repair by interacting with the human 3-methyladenine DNA glycosylase; Biochemistry, 2000, 39, 8032–8038. 97. Kartalou, M.; Essigmann, J.M.; Recognition of cisplatin adducts by cellular proteins; Mutation Res., 2001, 478, 1–21. 98. Wozniak, K.; Blasiak, J.; Recognition and repair of DNA-cisplatin adducts; Acta Biochim. Pol., 2002, 49, 583–596. 99. Bellon, S.F.; Coleman, J.H.; Lippard, S.J.; DNA unwinding produced by site-specific intrastrand cross- links of the antitumor drug cis-diamminedichloroplatinum(II); Biochemistry, 1991, 30, 8026–8035. 100. Bellon, S.F.; Lippard, S.J.; Bending studies of DNA site-specifically modified by cisplatin, trans-diamminedichloroplatinum(II) and cis-Pt(NH3)2(N3-cytosine)Cl+; Biophys. Chem., 1990, 35, 179–188. 101. Huang, J.C.; Zamble, D.B.; Reardon, J.T.; Lippard, S.J.; Sancar, A.; HMG-domain proteins specifically inhibit the repair of the major DNA adduct of the anticancer drug cisplatin by human excision nuclease; Proc. Natl. Acad. Sci. USA, 1994, 91, 10394–10398. 102. Plooy, A.C.M.; Fichtinger-Schepman, A.M.J.; Schutte, H.H.; van Dijk, M.; Lohman, P. H.M.; The quantitative detection of various Pt-DNA-adducts in chinese hamster ovary cells treated with cisplatin; application of immunochemical techniques; Carcinogenesis, 1985, 6, 561–566. 103. Dijt, F.J.; Fichtinger-Schepman, A.M.J.; Berends, F.; Reedijk, J.; Formation and repair of cisplatin-induced adducts to DNA in cultured normal and repair-deficient human fibroblasts; Cancer Res., 1988, 48, 6058–6062. 104. Fujiwara, Y.; Tatsumi, M.; Sasaki, M.S.; Cross-link repair in human cells and its possible defect in Fanconi’s anemia cells; J. Mol. Biol., 1977, 113, 635–649. 105. Calsou, P.; Frit, P.; Salles, B.; Repair synthesis by human cell extracts in cisplatin-damaged DNA is prefentially determined by minor adducts; Nucleic Acids Res., 1992, 20, 6363–6368. 106. Fisher, L.A.; Bessho, M.; Bessho, T.; Processing of a psoralen DNA interstrand cross-link by XPF-ERCC1 complex in vitro; J. Biol. Chem., 2008, 283, 1275–1281. 107. Chvalova, K.; Brabec, V.; Kasparkova, J.; Mechanism of the formation of DNA-protein cross-links by antitumor cisplatin; Nucleic Acids Res., 2007, 35, 1812–1821. 108. Olinski, R.; Wedrychovski, A.; Schmidt, W.N.; Briggs, R.C.; Hnilica, L.S.; In vivo DNAprotein cross-linking by cis- and trans-diamminedichloroplatinum(II); Cancer Res., 1987, 47, 201–205. 109. Plooy, A.C.M.; Van Dijk, M.; Lohman, P.H.M.; Induction and repair of DNA cross-links in Chinese hamster ovary cells treated with various platinum coordination compounds in relation to platinum binding to DNA, cytotoxicity, mutagenicity, and antitumor activity; Cancer Res., 1984, 44, 2043–2051. 110. Olinski, R.; Briggs, R.C.; DNA-protein cross-linking in L1210 cells and resistant to cisdiamminedichloroplatinum(II); Mol. Biol. Rep., 1991, 15, 81–86. 111. Richmond, T.J.; Davey, C.A.; The structure of DNA in the nucleosome core; Nature, 2003, 423, 145–150. 112. Wang, D.; Hara, R.; Singh, G.; Sancar, A.; Lippard, S.J.; Nucleotide excision repair from site-specifically platinum-modified nucleosomes; Biochemistry, 2003, 42, 6747–6753. 113. Reardon, J.T.; Vaisman, A.; Chaney, S.G.; Sancar, A.; Efficient nucleotide excision repair of cisplatin, oxaliplatin, and bis-aceto-ammine-dichloro-cyclohexylamine-platinum(IV) (JM216) platinum intrastrand DNA diadducts; Cancer Res., 1999, 59, 3968–3971. 114. Hartwig, J.F.; Lippard, S.J.; DNA binding properties of cis-Pt(NH3)(C6H11NH2)Cl2, a metabolite of an orally active platinum anticancer drug; J. Am. Chem. Soc., 1992, 114, 5646–5654. 115. Malina, J.; Novakova, O.; Vojtiskova, M.; Natile, G.; Brabec, V.; Conformation of DNA GG intrastrand cross-link of antitumor oxaliplatin and its enantiomeric analog; Biophys. J., 2007, 93, 3950–3962. 116. Malina, J.; Hofr, C.; Maresca, L.; Natile, G.; Brabec, V.; DNA interactions of antitumor cisplatin analogs containing enantiomeric amine ligands; Biophys. J., 2000, 78, 2008–2021.

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117. Malina, J.; Kasparkova, J.; Natile, G.; Brabec, V.; Recognition of major DNA adducts of enantiomeric cisplatin analogs by HMG box proteins and nucleotide excision repair of these adducts; Chem. Biol., 2002, 9, 629–638. 118. Delalande, O.; Malina, J.; Brabec, V.; Kozelka, J.; Chiral differentiation of DNA adducts formed by enantiomeric analogues of antitumor cisplatin is sequence-dependent; Biophys. J., 2005, 88, 4159–4169. 119. Reedijk, J.; Improved understanding in platinum antitumour chemistry; Chem. Commun., 1996, 801–806. 120. Farrell, N.; Current status of structure-activity relationships of platinum anticancer drugs: Activation of the trans geometry. In Metal Ions in Biological Systems, Vol. 32, Sigel, A. and Sigel, H.; Eds., Marcel Dekker, Inc., New York, Basel, Hong Kong, 1996, 603–639. 121. Coluccia, M.; Natile, G.; Trans-platinum complexes in cancer therapy; Anti-Cancer Agents Med. Chem., 2007, 7, 111–123. 122. Kasparkova, J.; Novakova, O.; Marini, V.; Najajreh, Y.; Gibson, D.; Perez, J.-M.; Brabec, V.; Activation of trans geometry in bifunctional mononuclear platinum complexes by a piperidine ligand: Mechanistic studies on antitumor action; J. Biol. Chem., 2003, 278, 47516–47525. 123. Marini, V.; Christofis, P.; Novakova, O.; Kasparkova, J.; Farrell, N.; Brabec, V.; Conformation, protein recognition and repair of DNA interstrand and intrastrand cross-links of antitumor trans-[PtCl2(NH3)(thiazole)]; Nucleic Acids Res., 2005, 33, 5819–5828. 124. Coluccia, M.; Nassii, F.; Loseto, F.; Boccarelli, A.; Marigió, M.A.; Giordano, D.; Intini, F.P.; Caputo, P.; Natile, G.; A trans-platinum complex showing higher antitumor activity than the cis congeners; J. Med. Chem., 1993, 36, 510–512. 125. Brabec, V.; Vrana, O.; Novakova, O.; Kleinwachter, V.; Intini, F.P.; Coluccia, M.; Natile, G.; DNA adducts of antitumor trans-[PtCl2(E-imino ether)2]; Nucleic Acids Res., 1996, 24, 336–341. 126. Novakova, O.; Kasparkova, J.; Malina, J.; Natile, G.; Brabec, V.; DNA-protein crosslinking by trans-[PtCl2(E-iminoether)2]. A concept for activation of the trans geometry in platinum antitumor complexes; Nucleic Acids Res., 2003, 31, 6450–6460. 127. Farrell, N.; Polynuclear platinum drugs. In Metal Ions in Biological Systems, Vol. 42, Sigel, A. and Sigel, H.; Eds., Marcel Dekker, Inc., New York, Basel, 2004, 251–296. 128. Perego, P.; Gatti, L.; Caserini, C.; Supino, R.; Colangelo, D.; Leone, R.; Spinelli, S.; Farrell, N.; Zunino, F.; The cellular basis of the efficacy of the trinuclear platinum complex BBR 3464 against cisplatin-resistant cells; J. Inorg. Biochem., 1999, 77, 59–64. 129. Roberts, J.D.; Beggiolin, G.; Manzotti, C.; Piazzoni, L.; Farrell, N.; Comparison of cytotoxicity and cellular accumulation of polynuclear platinum complexes in L1210 murine leukemia cell lines; J. Inorg. Biochem., 1999, 77, 47–50. 130. Farrell, N.; Polynuclear charged platinum compounds as a new class of anticancer agents: Toward a new paradigm. In Platinum-based Drugs in Cancer Therapy, Kelland, L.R. and Farrell, N.P.; Eds., Humana Press Inc, Totowa, NJ, 2000, 321–338. 131. Brabec, V.; Kasparkova, J.; Vrana, O.; Novakova, O.; Cox, J.W.; Qu, Y.; Farrell, N.; DNA modifications by a novel bifunctional trinuclear platinum Phase I anticancer agent; Biochemistry, 1999, 38, 6781–6790. 132. Fink, D.; Nebel, S.; Aebi, S.; Zheng, H.; Cenni, B.; Nehme, A.; Christen, R.D.; Howell, S.B.; The role of DNA mismatch repair in platinum drug resistance; Cancer Res., 1996, 56, 4881–4886. 133. Aebi, S.; Fink, D.; Gordon, R.; Kim, H.K.; Zheng, H.; Fink, J.L.; Howell, S.B.; Resistance to cytotoxic drugs in DNA mismatch repair-deficient cells; Clin. Cancer Res., 1997, 3, 1763–1767. 134. Johnson, S.W.; Ferry, K.V.; Hamilton, T.C.; Recent insights into platinum drug resistance in cancer; Drug Resist. Updates, 1998, 1, 243–254. 135. Drummond, J.T.; Anthoney, A.; Brown, R.; Modrich, P.; Cisplatin and adriamycin resistance are associated with MutL alpha and mismatch repair deficiency in an ovarian tumor cell line; J. Biol. Chem., 1996, 271, 19645–19648.

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136. Fink, D.; Nebel, S.; Aebi, S.; Nehme, A.; Howell, S.B.; Loss of DNA mismatch repair due to knockout of MSH2 or PMS2 results in resistance to cisplatin and carboplatin; Int. J. Oncol., 1997, 11, 539–542. 137. Ferry, K.V.; Fink, D.; Johnson, S.W.; Nebel, S.; Hamilton, T.C.; Howell, S.B.; Decreased cisplatin damage-dependent DNA synthesis in cellular extracts of mismatch repair deficient cells; Biochem. Pharmacol., 1999, 57, 861–867. 138. Calsou, P.; Frit, P.; Salles, B.; Double strand breaks in DNA inhibit nucleotide excision repair in vitro; J. Biol. Chem., 1996, 271, 27601–27607. 139. Frit, P.; Calsou, P.; Chen, D.J.; Salles, B.; Ku70/Ku80 protein complex inhibits the binding of nucleotide excision repair proteins on linear DNA in vitro; J. Mol. Biol., 1998, 284, 963–973. 140. Frit, P.; Canitrot, Y.; Muller, C.; Foray, N.; Calsou, P.; Marangoni, E.; Bourhis, J.; Salles, B.; Cross-resistance to ionizing radiation in a murine leukemic cell line resistant to cis-dic hlorodiammineplatinum(II): Role of Ku autoantigen; Mol. Pharmacol., 1999, 56, 141–146. 141. Bhattacharyya, A.; Ear, U.S.; Koller, B.H.; Weichselbaum, R.R.; Bishop, D.K.; The breast cancer susceptibility gene BRCA1 is required for subnuclear assembly of Rad51 and survival following treatment with the DNA cross-linking agent cisplatin; J. Biol. Chem., 2000, 275, 23899–23903. 142. Caldecott, K.; Jeggo, P.; Cross-sensitivity of gamma-ray-sensitive hamster mutants to cross-linking agents; Mutation Res., 1991, 255, 111–121. 143. De Silva, I.U.; McHugh, P.J.; Clingen, P.H.; Hartley, J.A.; Defects in interstrand cross-link uncoupling do not account for the extreme sensitivity of ERCC1 and XPF cells to cisplatin; Nucleic Acids Res., 2002, 30, 3848–3856. 144. Cummings, M.; Higginbottom, K.; McGurk, C.J.; Wong, O.G.-W., Koberle, B.; Oliver, R.T.D.; Masters, J.R.; XPA versus ERCC1 as chemosensitising agents to cisplatin and mitomycin C in prostate cancer cells: Role of ERCC1 in homologous recombination repair; Biochem. Pharmacol., 2006, 72, 166–175. 145. Selvakumaran, M.; Pisarcik, D.A.; Bao, R.; Yeung, A.T.; Hamilton, T.C.; Enhanced cisplatin cytotoxicity by disturbing the nucleotide excision repair pathway in ovarian cancer cell lines; Cancer Res., 2003, 63, 1311–1316. 146. Brabec, V.; Kasparkova, J.; Modifications of DNA by platinum complexes: Relation to resistance of tumors to platinum antitumor drugs; Drug Resist. Updates, 2005, 8, 131–146. 147. Jung, Y.; Lippard, S.J.; Direct cellular responses to platinum-induced DNA damage; Chem. Rev., 2007, 107, 1387–1407. 148. Teletchéa, S.; Komeda, S.; Teuben, J.-M., Elizondo-Riojas, M.A.; Reedijk, J.; Kozelka, J.; A pyrazolato-bridged dinuclear platinum(II) complex induces only minor distortions upon DNA-binding; Chem. Eur. J., 2006, 12, 3741–3753. 149. Komeda, S.; Lutz, M.; Spek, A.L.; Yamanaka, Y.; Sato, T.; Chikuma, M.; Reedijk, J.; A novel isomerization on interaction of antitumor-active azole-bridged dinuclear platinum(II) complexes with 9-ethylguanine. Platinum(II) atom migration from N2 to N3 on 1,2,3-triazole; J. Am. Chem. Soc., 2002, 124, 4738–4746. 150. Komeda, S.; Kalayda, G.V.; Lutz, M.; Spek, A.L.; Yamanaka, Y.; Sato, T.; Chikuma, M.; Reedijk, J.; New isomeric azine-bridged dinuclear platinum(II) complexes circumvent cross-resistance to cisplatin; J. Med. Chem., 2003, 46, 1210–1219. 151. Komeda, S.; Bombard, S.; Perrier, S.; Reedijk, J.; Kozelka, J.F.; Kinetic study of azolebridged dinuclear platinum(II) complexes reacting with a hairpin-stabilized doublestranded oligonucleotide; J. Inorg. Biochem., 2003, 96, 357–366. 152. Spiegel, K.; Magistrato, A.; Carloni, P.; Reedijk, J.; Klein, M.L.; Azole-bridged diplatinum anticancer compounds. Modulating DNA flexibility to escape repair mechanism and avoid cross resistance; J. Phys. Chem. B, 2007, 111, 11873–11876. 153. Madhusudan, S.; Hickson, I.D.; DNA repair inhibition: a selective tumour targeting strategy; Trends Mol. Med., 2005, 11, 503–511. 154. Hubscher, U.; Maga, G.; Spadari, S.; Eukaryotic DNA polymerases; Annu. Rev. Biochem., 2002, 71, 133–163.

7 Telomeres and Telomerase: Potential Targets for Platinum Complexes Isabelle Ourliac-Garnier, Razan Charif and Sophie Bombard

7.1 Function of Telomeres Telomeres are nucleoprotein structures that protect the very end of linear chromosomes. Telomeric DNA consists of tandemly repeated G-rich sequences of DNA, which differ from one species to another, being (TTAGGG)n in humans. These sequences of about 4–10 kbp protrude at the 3′ extremity by a single strand of the G-rich sequence of about 50 to 200 bases. Firstly, telomeres preserve the stability and the structural integrity of chromosomes;1–4 indeed, telomere dysfunction leads to chromosome instability and abnormalities, senescence or apoptosis, and represents potentially the most widespread cause of genome instability in cancer.5 So, concretely, telomeres avoid end-to-end fusion of chromosomes by preventing the chromosomes ends from being recognised as DNA double-strand breaks (DSB) and consequently from being repaired.6 Secondly, telomeres preserve the genome integrity by protecting chromosomes from a loss of genetic information due to the ‘end replication problem’: a progressive erosion of the 5′ extremity of each chromosome that happens at each round of replication.7–10 Indeed, DNA replication involves RNA primers, named Okazaki fragments, which are necessary for DNA synthesis from the 5′ to the 3′ extremities. The last replication fork is supposed to be subtelomeric, consequently, telomeric DNA is replicated unidirectionally and the G-rich strand is synthesized discontinuously, whereas the C-rich one is replicated continuously (Figure 7.1). Then, conventional DNA polymerases replace Okazaki Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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G C

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exonuclease 3'

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Figure 7.1 End replication of telomeres and processing. The G-rich strand is replicated discontinuously, whereas the C-rich one is replicated continuously. The RNA Okazaki fragments (in dark grey) at the 5′ end of the daughter DNA (dotted lines) is not replaced by conventional DNA polymerases, the parental G-rich strand is consequently not completely replicated, leading to the G-overhang formation. Moreover, the 5′ end of the C-rich strand of telomeres (daughter or parental strand) can also be digested by a 5′ to 3′ nuclease which contributes to the formation of the G-overhang

fragments by DNA, except at the 5′ end of the daughter DNA, which becomes shorter. Consequently, the parental G-rich strand is not completely duplicated leading to the formation of a G-rich overhang. Moreover, the telomere shortening mechanism is also due to a particular processing of the C-rich strand.11,12 Particularly, it has been proposed that Apollo, a protein involved in DNA repair and colocalised with telomeric proteins, contributes to the resection that occurs at the 5′ C-rich strand thanks to its 5′-exonuclease activity.13 Consequently, in normal cells, telomeres erode progressively at each round of replication and can shorten until a length limit (Hayflick limit) that induces a signal in the cell, leading to senescence.14 Telomeres are therefore considered as a mitotic clock that allows the cell to divide a limited number of times before entering senescence.15,16

7.2 Structure of Telomeres Telomeres have a well-organised structure comprising telomeric DNA and a package of specific proteins that are called the telosome17 or the shelterin18 and cap the chromosome extremity. The shelterin is composed of six proteins: TRF1, TRF2, POT1, TIN2, TPP1 and Rap1. Three of these are specifically and directly bound to

Structure of Telomeres TPP1

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Figure 7.2 The shelterin complex and associated proteins. T-loop formation has been proposed as a structure capping the end of telomeres. It consists of the invasion of the 3′ Goverhang into the duplex telomeric DNA. Associated telomeric proteins are involved in this complex: TRF1, TRF2 and POT1 bind directly to telomeric DNA, whereas TPP1, TIN2 and RAP1 interact with them. Other proteins that are not specific for telomeres have also been associated with this complex. They include Apollo, which appears to protect chromosome ends from being processed as DNA damage, and DNA damage proteins: PINX1, Ku 70/80, TANK1/2, ATM-kinase, PARP2, ERCC1/XPC, MRE11, Rad 50, Rbs1

telomeric DNA: TRF1 and TRF2 bind to the double-stranded structure through their Myb-type DNA binding domain19–21 and POT1 binds to the G-rich 3′ overhang through two oligonucleotide/oligosaccharide binding domains.22 The three other proteins allow the connection between the double-stranded telomeric DNA and its 3′ extremity: TIN2 binds to TRF1 and TRF2; TPP1 binds to TIN2 and POT1; Rap1 binds to TRF2.23 Two telomeric proteins, TRF2 and POT1 are directly involved in the maintenance of the structure of telomeres.24 In particular, as shown by an electron microscopy study, TRF2 is thought to promote the formation of the t-loop, which consists of an invasion of the G-rich overhang in the double-stranded telomeric DNA,25 and is supposed to protect the chromosome extremity from degradation26 (Figure 7.2). Moreover, it has been found that expression of a dominant-negative mutant allele of TRF2 induces: (i) recognition of telomeres as DSB, (ii) activation of the ATM/p53 DNA damage response pathway, (iii) end-to-end fusions of chromosomes and (iv) accelerated senescence entry.27–29 TRF2 is overexpressed in some cancer cell lines,30–32 and it has been shown that the inhibition of TRF2 can reduce tumourigenicity of melanoma cancer cell lines.33 This suggests that TRF2 is strongly involved in telomere capping in cancer cells and could be a good target for anticancer therapy. POT1 has been shown to act either as an activator or as a repressor of the enzyme responsible of telomere elongation, telomerase (see below), via its interaction with TRF1 and TPP1.34–38 Dysfunctional or uncapped telomeres, created through destabilization of the shelterin by TRF2 or POT1 inhibition, leads to the DNA damage response,3,35,39–42 loss of the 3′-G-overhang and aberrant homologous recombination.43,44 In conclusion, the protective function of telomeres depends essentially on the stability and integrity of the shelterin, which could be perturbed by modification of the amount of telomeric proteins bound to DNA.

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7.3 Telomerase Cells escape limits of proliferation by maintaining a constant length of their telomeres. This is mainly due to telomerase, a telomere-specific reverse transcriptase that was first identified in 1985.45 Telomerase is activated and expressed in 85% of cancer cells, in germ cells, some stem cells46,47 and at a very low rate in some epithelial and intestinal cells.48 It functions by adding telomeric repeats GGTTAG at the end of the 3′-G-overhang of telomeres, using an RNA template complementary to the telomeric sequence.49 This mechanism prevents replicative senescence, thus conferring on cells unlimited proliferation.50 Indeed, it has been shown that when telomerase was introduced into normal cells, it resulted in an extension of their life span.51 Therefore, telomerase has been proposed as one of the hallmarks of cancer cells52 and as a cancer marker or prognosis factor.53 Telomerase consists of two main components: a protein component reverse transcriptase, hTERT, (127 kD) that contains conserved catalytic reverse transcriptase motifs,54 and a large RNA of 451 nucleotides, hTR, (153kD). It contains a template complementary to telomeric sequences, and other critical conserved regions essential for the assemblage of hTERT with hTR or for telomerase activity55,56 (Figure 7.3). Recently, a third partner, dyskerin, a pseudouridine synthase belonging to the H/ACA box ribonucleoproteins, has been identified as an essential component of the telomerase complex57,58 that probably binds to the H/ACA motif of hTR. Moreover the telomerase trimeric complex has been proposed to be associated in cells with about 30 other regulatory proteins involved in biogenesis, trafficking and recruitment to telomeres.57,59 hTR

hTR

CAAUCCCAAUC TTAGGGTTAGG

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Figure 7.3 Telomerase is composed of two main components: a protein component that contains the catalytic reverse transcriptase motif (hTERT) and a large RNA of 451 nucleotides (hTR) containing the template region able to elongate the 3′ extremity of telomeres. Other proteins, that are not necessary for the in vitro telomerase activity, have been associated with telomerase in vivo, such as dyskerin. In addition to the template, many structural domains of hTR are conserved among the species (in grey boxes). They are involved in the assemblage of hTR with hTERT, in telomerase activity or in hTR processing

G-Quadruplex Structures

213

Since telomerase activity is elevated in the vast majority of tumours, telomerase participates in the immortality of cells, thanks to telomere elongation. Since telomeres are critically shortened in tumours versus normal tissues, it is thus conceivable to inhibit telomerase as a strategy for cancer therapy. Inhibition of telomerase has been performed using oligonucleotides targeting hTR or the mRNA of hTERT, or nucleoside analogues and nonnucleoside inhibitors targeting the catalytic centre of hTERT. They have been shown to induce progressive telomere shortening and senescence or apoptosis of treated cancer cells.60–63

7.4 G-Quadruplex Structures and Small Molecules It has been recognised that the 3′-G-overhang of the telomere, (TTAGGG)n, is able to fold, in vitro, into G-quadruplex structures. The structures consist of G-quartets that are stacked in a planar arrangement of four guanines associated by Hoogsteentype hydrogen bonds. Four consecutive repeats of guanines in the telomeric sequence are required for its folding in an intramolecular G-quadruplex structure. The monovalent cations Na+ or K+ are required to stabilise these structures by interacting with the eight carbonyl oxygens of two adjacent G-quartets64,65 (Figure 7.4A). Although there is no direct proof of G-quadruplexes in vivo, recent biological evidence is in favour of their existence in the ciliate Stylonychia66 and in humans.67 Moreover, some proteins have been shown to promote their formation68 and to induce their unfolding.69 G-quadruplexes can also be formed outside the telomere region, because intramolecular G-quadruplex-forming sequence motifs are prevalent in the genome,70,71 and particularly enriched in gene promoters.72 Moreover, it has been shown that gene expression depending on these promoters could be modulated by G-quadruplex binders.73 Crystallographic and NMR studies of human telomeric quadruplex structures have revealed a high degree of polymorphism in the structures, depending on the folding of the DNA backbone, the orientation of the strands (parallel or antiparallel) and the conformation of the TTA loops (reverse, lateral or transversal). Four main G-quadruplex structures have been described for telomeric DNA: parallel,74 antiparallel75 and two mixed-hybrid structures76,77 (Figures 7.4B–E). Each structure has been determined in the presence of a particular monovalent cation (Na+ or K+), but it is now clear that these structures coexist in solution, whatever the cation.78–81 Since the substrate of telomerase is the linear extremity of the 3′-G-overhang of telomeric DNA, it has been shown that its folding in G-quadruplex structures may impede its recognition by telomerase and may interfere with telomere elongation.82 Therefore, the stabilization of quadruplex DNA structures by small molecules has been proposed as a new strategy in order to inhibit telomerase and interfere with telomere maintenance in tumour cells.53 Many molecules have been synthesised in order to target G-quadruplexes and then tested on telomerase activity and cell proliferation. The paradigm for telomerase inhibitors is that senescence of cancer cells will occur after many rounds of replication because of progressive telomere shortening. Some of the G-quadruplex binders have been shown to follow this rule.83 However, the test used

Telomeres and Telomerase

214

A

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Figure 7.4 Schematic structures of the G-quadruplexes formed by the human telomeric sequence: (A) the G-quartet consists of four guanines linked through Hoogsteen-type hydrogen bonds; (B) d[AGGG(TTAGGG)3] antiparallel, Na+ solution;75 (C) d[AGGG(TTAGGG)3] parallel, K+ (crystal); (D) and (E) d[TAGGG(TTAGGG)3])77 and d[TAGGG(TTAGGG)3TT],76 K+ solution, mixed-hybrids. Syn and anti guanines are coloured white and grey, respectively

G-Quadruplex Structures

215

Figure 7.5 Chemical formulae of G-quadruplex ligands that inhibit cell proliferation in a way independent of telomerase elongation

for appreciation of in vitro telomerase inhibition, TRAP (telomeric repeat amplification protocol) has been revaluated. The results indicate that the inhibitory effect of many G-quadruplex binders on telomerase elongation has been overestimated because these molecules were shown to be good inhibitors of the PCR step used for the amplification protocol.84 Moreover, some of these G-quadruplex binders (telomestatin, RHPS4, BRACO 19) (Figure 7.5) have been shown to inhibit cell proliferation in a way independent of telomerase elongation.85–90 It has been suggested that they stabilize the multiple G-quadruplex structures that can be formed transiently along telomeric DNA,91,92 thus altering the recognition of the telomeric sequences by proteins from the shelterin (TRF2 and POT1)85–89 or that they can inhibit telomere replication.93 The resulting increase in genomic instability has been found to depend on the type of cancerous cells.33,90 This selective toxicity for cancer cells can be explained by the fact that telomeres from normal and cancer cells exhibit differences in structure, stability and accessibility.94 There is therefore a growing interest in considering G-quadruplexes as potential targets for new antitumour drugs.64,65,91–93,95,96 The past few years have seen considerable development in design and synthesis of novel and efficient G-quadruplex binders.97 At present, the most efficient one is telomestatin.85–88,98–100 In general, an effective G-quadruplex binder is planar and, in most of cases, positively charged. On the basis of the very few structural data of G-quadruplex binders available nowadays, it has been proposed that the interaction occurs via π-stacking and electrostatic interactions.101–106 Design of new binders that are easily synthesized is still a challenge. A promising recent approach relies on the use of metallo-organic complexes, based on the assumption that the central metal could be positioned over the cation channel of the quadruplex, thereby optimizing the stacking interactions with the

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Figure 7.6 Chemical formulae of selected metallo-organic G-quadruplex ligands

external G-quartet.107 Therefore, transition metals are considered very appealing, thanks to their multiple and controllable geometries, the possible modifications of their coordination environment and their easy synthetic access. Moreover, it has been highlighted that the metal centre is a key parameter governing the recognition of G-quadruplex using a family of terpyridine metallo-organic complexes.108 The pertinence of this approach has been demonstrated by the high selectivity of saphen (Ni(II)),109 terpyridine (Cu(II) and Pt(II)),108 porphyrins (Mn(III),110 Cu(II)111,112 and Ni(II)113, 114) and phthalocyanine derivatives (Zn(II) and Ni(II)115,116) for Gquadruplexes (Figure 7.6).

7.5 Cisplatin The antitumour effect of the drug cisplatin, cis-[Pt(II)Cl2(NH3)2], was discovered in 1965.117 The cellular target of cisplatin, which triggers its anticancer activity, is

Interactions of Platinum Complexes with G-Quadruplex Structures 217

assumed to be DNA. Cisplatin becomes activated intracellularly by the aquation of the two chloride leaving groups, leading to cis-[Pt(II)(NH3)2(H2O)2]2+, and subsequently covalently binds to DNA, mainly forming intrastrand crosslinks on DNA between two adjacent guanines d(GpG) (65%) or adjacent adenine and guanine, d(ApG) (25%), by coordinating to N7 of purine bases.118–120 The antitumour effect of cisplatin is attributed to the formation of the major GG adduct because the tumour response correlates with the level of GG adducts. The recognition and processing by DNA damage-response proteins120 induce signal transduction pathways that lead to apoptosis or necrosis. However, although cellular responses to platinum-induced damage have been widely studied119, 121 all the biological targets are not fully revealed. Since human telomeres consist of long tandem repeats of TTAGGG sequences, they are potential targets for platinum complexes. Given that telomeric nucleosomes are intrinsically more mobile under physiological conditions than nucleosomes formed on an average DNA sequence,94 one can envisage that platination of telomeres could be kinetically favoured over average DNA. It is thus conceivable that cisplatin may exert part of its anticancer activity via the formation of DNA adducts at telomeres, altering the telomere structure and thus impeding its access to telomerase, and/or lead to a loss of specific telomere binding proteins. It is also possible that cisplatin could influence directly the telomerase activity by binding to its RNA core, hTR. This review will present the advances in this field.

7.6 Interaction of Cisplatin and Related Platinum Complexes with G-Quadruplex Structures Since metallo-organic complexes appeared recently as promising new G-quadruplex binders,97 square planar Pt(II) complexes with extended aromatic ligands have been prepared. Indeed, they provide a p-surface that is compatible with the G-quartet motif. Although some of them have been extensively studied as duplex DNA binders, they have been less studied for their recognition of G-quadruplex DNA. Only a few examples of platinum–organic G-quadruplex binders are known to date. They belong to two different classes: (i) Pt(II) complexes without any labile ligand, unable to undergo platination reaction on the G-quadruplexes and therefore considered as reversible binders; (ii) mono-functional Pt(II) complexes containing one labile ligand (Cl−) potentially able to covalently link G-quadruplexes and considered as potential irreversible binders in addition to their stabilizing effect. (i) Bipyridine and phenanthroimidazole ethylenediamine platinum (II) complexes (Figure 7.7A) have been shown to interact with a 12-mer intermolecular quadruplex-forming sequence (T4G4T4)4 with two platinum complexes per quadruplex.122 Binding constants are in the range 106–107 M−1 increasing with the p-surface of the complexes. They induce significant stabilization of the intermolecular G-quadruplex structure and increased preference of two orders of magnitude for quadruplex over duplex. This binding mode is consistent with an end-stacking platinum complex located at each side of the quadruplex structure. Molecular modelling studies of the Pt(II) complexes with the quadruplex

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Figure 7.7 Chemical formulae of selected platinum–organic G-quadruplex ligands known to stabilize specifically G-quadruplexes over duplex DNA. Platinum complexes (A) are unable to platinate G-quadruplexes once bound to it, whereas platinum complexes (B) are potentially able to platinate G-quadruplexes since they are monofunctional. However, this possibility has not been yet investigated

structure of (T4G4T4)4 showed p-stacking interaction of the aromatic system with the guanines of both external G-quartets. Surprisingly, the Pt(II) is offcentre due to the presence of three hydrogen bonds between the ethylenediamine ligand and the DNA phosphate backbone.122 (ii) A mono-substituted phenanthroline has been synthesized and used as a tridendate N,N,N-ligand to coordinate to Pt(II)123 (Figure 7.7B). This design was based on molecular modelling suggesting that the Pt(II) ion could be positioned over the cation channel at the centre of an accessible G-quartet, optimizing the stacking interactions. In the same context, three Pt(II)–terpyridine complexes have been synthesized (Figure 7.7B).108 The monosubstituted phenanthroline– Pt(II) complex induces a high degree of stabilization for quadruplexes (increase in the melting temperature over 20 °C) and is selective for quadruplex structure over duplex by a factor of 40. A comparable increase in melting temperature (11–16 °C) was observed with the Pt(II)–terpyridine complexes. Moreover, the metal-free ligand has considerably lower effect on the stabilization than Pt(II) complexes, suggesting that the presence of a Pt(II) centre plays an important role in quadruplex stabilization. Since both platinum complexes are monofunctional, (one Cl− labile ligand), they should be able to link covalently to purines of G-quadruplexes. However, this possibility has not yet been investigated by the authors. Studies from our group pointed out that G-quadruplex structures, against all expectations, were potential targets for Pt(II) coordination.78,124–127). We first analysed the possible platination of two quadruplex structures of the Tetrahymena

Interactions of Platinum Complexes with G-Quadruplex Structures 219 5'

T14

G3

G15

G4

T14

5'

T13 G3

G12

T13 G15

G12

3' G24

G21

G16 G5

G16

T20

G10

G6

G17

G22

G17

G23

G5 T19

G22

G21

G9 T7 G18

T8

T19

G11

G4

G11

G23

G10 G6 G9 T7

G24

G18 T8

3'

T20 A

B

Figure 7.8 Schematic structures of the G-quadruplex formed by the telomeric sequence of Tetrahymena. The antiparallel structure (A) has been solved by chemical probes128 and the mixed-hybrid one (B) has been solved by NMR130

sequence, (T2G4)4, which have been structurally determined by Williamson et al.128, Henderson et al.129 and Wang and Patel.130 These structures contain four guanines in their loops in addition of the three G-quartets (Figure 7.8). The four guanines of a G-quartet have their N7 atom involved in hydrogen bonding. Therefore, within a G-quadruplex structure, we postulated that only the four guanines not included in the G-quartets should react with Pt(II) complexes. The two quadruplex structures exhibit various loop and strand orientations (Figure 7.8), in which different guanines, not included in the G-quartets, are exposed: G3, G9, G15, G21 in the antiparallel structure (Figure 7.8A) and G6, G9, G15, G24 in the hybride structure (Figure 7.8B). Using the monofunctional triammine complex [Pt(II)(NH3)3(H2O)]2+, we have shown that only the four guanines G3, G9 G15 and G21, (but not G6 and G24) are platinated, suggesting the existence of the antiparallel structure as the major one in solution. However, the four guanines were not equally accessible for platination. Molecular dynamics simulations were used in order to interpret these results and allowed us to conclude that the relative accessibility of the N7 atom of the free guanines depends on their stacking with the neighbouring bases and on their binding with the central cation, positioned within the loops.124 Therefore [Pt(II)(NH3)3(H2O)]2+ could be considered as a useful tool to check the accessibility of the nucleophilic sites of purines. Furthermore, the difunctional Pt(II) complexes, cis- and trans[Pt(II)(NH3)2(H2O)2]2+, were shown to crosslink both ends of the quadruplex structure of (T2G4)4, since G3–G15 and G9–G21 crosslinks were formed. Molecular dynamics simulations indicated that these crosslinks occurred without perturbing the overall structure and the quartet stacking.124

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It was therefore challenging to determine if such Pt(II) complexes were able to cross link the quadruplex structures of the human telomeric sequence, AG3(T2AG3)3. Indeed, they contain only four adenines in the loops and stem that are accessible to Pt(II) complexes (Figure 7.3). Moreover it was of pharmacological interest to establish if such a crosslink might exist and have similar cellular effects to those of compounds stabilizing the G-quadruplex structures.96,131 The results of the in vitro platination of the human G-quadruplex structures by the monofunctional platinum complex [Pt(II)(NH3)3(H2O)]2+ was surprising because, in addition to the four adenines, some guanines were platinated, whereas their N7 atom was expected to be involved in hydrogen bonds into the G-quartets. Thanks to unconstrained molecular dynamics simulations run on the antiparallel and on the parallel structures (Figure 7.9), it was possible to rationalize the platination of these specific guanines. Indeed, the simulations clearly showed that both structures were flexible and that the G-quartet located at the transversal loop side of the antiparallel structure could be disrupted, allowing the N7 atoms of G2, G10, G14 and G22 to be platinated. Moreover, two guanines located in the G-quartet of the 5′ extremity of the parallel structure were also shown to slip out of the G-quartet, rendering their N7 atoms (G8 and G20) accessible to platination. Since the adenines of the loops and some guanines of the G-quartets are potential platination sites, crosslinks of human telomeric G-quadruplex structures by difunctional Pt(II) complexes may be envisioned. Actually, we showed that the antiparallel structure was crosslinked by the difunctional Pt(II) complexes, cis- and trans-[Pt(II)(NH3)2(H2O)2]2+, between an adenine and a guanine, (A1–G10, A13–G22), as well as the two adenines located at both ends of the structure (A1–A13, A7–A19).125 These results indicate that purines of the parallel structure (A–A or A–G) are not close enough to be crosslinked by the mononuclear difunctional platinum

G2

G20

G10 G14

G-quartets

G8

G22

Anti-parallel structure Parallel structure of the human telomeric DNA sequence AG3(T2AG3)3

Figure 7.9 Conformation of the antiparallel and the parallel structures AG3(T2AG3)3 from molecular dynamics simulations. The top G-quartet of the anti-parallel structure can be completely disrupted whereas the top G-quartet of the parallel structure can lose two guanines. The slipping out of these guanines render their N7 atom accessible to platinum complexes. (Adapted from Biochemistry, 2005, 44, 10620–10634.)

Interactions of Platinum Complexes with G-Quadruplex Structures 221

complexes. In order to try to crosslink the parallel structure, we used the dinuclear difunctional platinum complexes, [{trans-PtCl(NH3)2}2H2N(CH2)nNH2]Cl2 (n = 2 or 6), in which two platinum coordination units are linked by a flexible diamine linker that allows long range crosslinks on duplex DNA.132 We found that the parallel structure was mainly crosslinked between two guanines at the 5′-end G-quartet (G8–G10) and (G2–G20), respectively, and that the antiparallel structure was crosslinked between two guanines at the 5′-end G-quartet (G2–G10), (G10–G22) and (G2–G14).78 All these results highlight that human G-quadruplex structures may serve as favourable platination targets, since some of their guanines can reversibly leave the G-quartet plane, thus rendering their N7 atom accessible to platinum complexes. Moreover, we have showed that platination of human telomeric G-quadruplexes by cisplatin was kinetically favoured in vitro by a factor of about two over a duplex containing a unique GG platination site.126 Based on these results we have designed new platinum complexes combining a G-quadruplex binder133 with Pt(II) in order to target more specifically Gquadruplex structures. Firstly, a platinum–quinacridine complex (Pt-MPQ) has been specifically designed to bind covalently in order to stabilize G-quadruplex structures.127 The platinum moiety can reach the opposite external G-quartet and bind to an accessible purine (Figure 7.10). Actually, this novel platinum hybrid complex is shown to trap preferentially the antiparallel quadruplex structure by linking to the guanines of the 5′-end G-quartet and also enhance quadruplex versus duplexDNA selectivity. More importantly, Pt-MPQ displays cytotoxic activities analogous to that of cisplatin on various cell lines. Another monofunctional hybrid platinum complex, a Pt(II)-acridine derivative, Pt-ACRAMTU, (Figure 7.10) has also been synthesized. Initially, it was known to form adenine adducts in duplex DNA. Later, it has been shown to bind preferentially to the adenines of the TTA loops of the G-quadruplex of human telomeric sequences and to react significantly faster with G-quadruplex than with duplex DNA.134 It was proposed that the acridine was stacked on one of the external G-quartets, which will favour the platination of an adenine of the neighbouring TTA loops. Interestingly, the two hybrid platinum complexes (Pt-MPQ and Pt-ACRAMTU) show major differences in their binding mechanisms to Gquadruplexes. It is conceivable that the length of the linker between the platinum moiety and the G-quadruplex binder governs the site of platination. However, the reversible stacking reaction is quicker than the irreversible platination reaction in both cases, suggesting that it is the binding reaction that will govern the platination site. For Pt-ACRAMTU, the binding of the acridine and the platination reaction will occur at the same side of the G-quadruplex. In this case, the guanines of the G-quartet are protected from platination due to the p-stacking interaction with the acridine. Thus, only N7 and N1 atoms of adenines in the loops are free and susceptible to interact with Pt(II). For Pt-MPQ, the quinacridine moiety binds at one side of the G-quadruplex and platination can occur at the opposite external G-quartet, thanks to the linker. Since the guanines of the 5′-end G-quartet of the antiparallel structure have been platinated by Pt-MPQ, it suggests that the quinacridine moiety binds at the opposite 3′-end G-quartet. Moreover, it should be noted that no adenine

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Pt 5'

A13

G10

G22

A1 G14

G2

G21

G9 G3

G15

G8 G4

G20 A19

G16

T18

T5 T12 5'

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A13 Pt

3' G22

A1 G14

G2

G21

G9 G15

G3 G8 G4

G20 G16

T5

T6

T17

A19 T18

Figure 7.10 Chemical formulae of two platinum–organic G-quadruplex binders that have been shown to stabilize selectively and to bind covalently to G-quadruplexes. The proposed binding mode on the antiparallel human telomeric G-quadruplex is indicated. For Pt-MPQ, the binding and platination sites are located at both sides of the G-quadruplex,127 whereas, for Pt-ACRAMTU,134 both sites are located at the same side of the G-quadruplex. Platination sites are represented by circles

was found to be platinated, indicating that the guanines, even when they are thought to be involved in the G-quartet, react more rapidly than adenines that are about ten times less nucleophilic.135 It is worth noticing that binding of the quinacridine moiety on the 5′-end G-quartet cannot be excluded, but it would prevent the platination of the G-quadruplex, since no platinable guanines are available on the opposite 3′-end.78 Since the four adenines and some guanines of the flexible G-quartets of the human G-quadruplex structure are efficiently platinated, the design of new platinum–hybrid compounds using different G-quadruplex binding motifs, represents an exciting challenge. Preliminary studies with these conjugates have revealed interesting biological responses worth further investigation.

Interactions of Platinum Complexes with Telomerase 223

7.7 Interaction of Cisplatin and Related Platinum Complexes with Telomeric DNA Duplexes Due to the presence of triple runs of guanines, a telomeric DNA duplex is a potential target for cisplatin. Firstly, it has been shown that the platinum content of a plasmid containing a 800bp (TTAGGG)n sequence is higher than in the remaining plasmid sequence, in proportion to the greater number of guanines.136 Secondly, it has been shown that longer telomeres are associated with resistance to cisplatin in melanoma cells.137 It is thus conceivable that the telomeric DNA–cisplatin interaction could be relevant for drug sensitivity/resistance status depending on the lengths of the telomere. However, it is worth noticing that this hypothesis has been invalidated in other cell lines.138 Thirdly, modified guanines are critical for association of telomere binding proteins since it has been shown that the presence of single or multiple 8-oxo-G lesions on telomeric sequences interfere with the in vitro binding of TRF1 and TRF2.139 All these results enable us to hypothesize that the modification of DNA by cisplatin could occur at telomeres and thus disturb the recognition of telomeric DNA by TRF1 and TRF2. Based on this hypothesis, we analysed the in vitro binding of TRF1 and TRF2 on the telomeric sequences (T2AG3)4/(C3TA2)4 platinated at a defined GGG site by cisplatin. We showed that the presence of cisplatin decreases the affinity of TRF2 more dramatically than that of TRF1 for these sequences.140 These results suggest that if there is any platination of telomeres by cisplatin in cancerous cells, it could have some impact on the binding of TRF2 and consequently could affect the integrity of the telomeres.

7.8 Interaction of Cisplatin and Related Platinum Complexes with Telomerase The RNA component of telomerase, hTR, is necessary for the reverse transcriptase activity of the enzyme. The template and the conserved structured regions (loops and pseudo-knot) of hTR are accessible binding sites,141 consequently hTR is a potential target for antitelomerase drugs. Antisense oligonucleotides, peptide nucleic acids (PNAs) directed against the template142 and small molecules directed against the tertiary structures143,144 have been shown to inhibit efficiently telomerase activity. The most promising analogue is GRN163L, a lipid conjugate to thiophosphoramidate145 that is used in breast cancer therapy.146 Since electrophilic agents could interact with the accessible nucleobases of hTR, the effect of cisplatin and some Pt(II) derivatives has been evaluated in vitro on telomerase activity using the TRAP assay.147 Particularly, novel platinum complexes bearing aromatic amines as bulky carrier groups have been prepared for this purpose.148,149 Inhibition of telomerase, dependent on Pt(II) complex interactions, has been observed, indicating platination of hTR; however, we can not exclude that the effect may be due to platination of essential nucleophilic amino acids of the proteic unit, hTERT. It is know that TRAP assay occurs with cells extracts during 30 minutes of incubation, followed by PCR amplification of the telomeric repeats.150 However, this incubation timescale is

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insufficient to allow total hydrolysis of the platinum complexes;151 indeed, the half life of hydrolysis of cisplatin and carboplatin is about 2 and 286 h, respectively. Therefore, the in vitro assay of telomerase activity is not fully informative, since a part of the platinum complexes remains inactive. Many workers have also investigated the cellular effects of cisplatin on telomere lengths, and telomerase residual activity and expression. However, the three parameters have never been taken into account at the same time. Until now, the effect of cisplatin on telomeres has not been well clarified. Concerning the effect of cisplatin on telomere length, the first study showed that telomere loss in Hela-treated cells (20 kbp in telomere length) was dependent on cisplatin dose. Low concentration of cisplatin (0.5 mM) induces telomere shortening after 24 h of treatment, whereas higher concentrations that blocked DNA replication in the S phase did not have any effect on telomere length.152 Cisplatin was also shown to induce the shortening of telomeres in hepatoma cells after 24 to 72 h treatment (0.8 to 50 mM).153 Since the rapid loss in telomere length was observed after only one to three rounds of replications, it probably did not result from telomerase inhibition, which induces a progressive shortening of telomeres.83 It can thus be proposed that platination of telomeres may inhibit telomere replication and/or could uncap the telomere, as already suggested for rapid loss of telomeres from quadruplex-binder-treated cells.93 A gradual shortening of telomeres was observed when yeast NER (nucleotide excision repair) mutant cells were treated by doses of cisplatin that did not affect cell viability.154 This suggests that the NER pathway, which is involved in repair of cisplatin adducts,119,155,156 may play a critical role in the repair and maintenance of telomeres. However, in another recent study it was found that cisplatin treatment (0.5–500 mM) induced no loss of telomeres after 2 to 72 h in three cell lines, independent of their initial telomere length (4–80 kbp) (neuroblastoma, Hela, acute lymphoblastic T cell).138 The results from these different studies were not conclusive and gave only an indication of the possibility that cisplatin could interfere with telomeres. It could depend on the susceptibility of the various cell lines used in the studies toward cisplatin. Moreover, the high cytotoxic activity of cisplatin makes it difficult to discriminate between overall cellular effects and the specific effect on telomerase and/or telomeres. Telomerase activity of cisplatin-treated cells has also been investigated. Telomerase expression (hTERT, mRNA or hTR quantification) has been controlled in parallel, in some cases, in order to find out if the decrease in telomerase activity was due to its downregulation or to its inactivation. Since the evaluation of this activity has been performed with TRAP assay150 after cell extraction, telomerase inhibition can reflect only irreversible inactivation of telomerase. This inactivation, which occurs upon cell treatment, could be attributed to covalent binding of cisplatin to telomerase (hTR or hTERT), but there is no proof of this event at this time. The results are still the subject of controversy. Upon cisplatin treatment, a decrease in telomerase activity has been found in human testicular cancer cells,157 associated with a downregulation of hTR. A decrease in telomerase activity has also been observed in hepatoma cells,153 breast cancer cells in culture and in xenograph tumours.158 In contrast, no decrease of telomerase activity has been detected after

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225

treatment by cisplatin of haematopoietic cancer cells when used at concentrations resulting in cytostatic growth inhibition159 in human ovarian adenocarcinoma cell lines,160 neuroblastoma, Hela and acute lymphoblastic T cells138 and of nasopharyngeal cancer cells.161 No significant downexpression of hTERT has been observed in MCF-7 cells treated with cisplatin.162 Since telomerase plays a role in cellular resistance to apoptosis, which is the primary mode of cell death induced by several drugs, combining telomerase inhibition with chemotherapeutic treatment may prove more efficient than each approach on its own. Many studies have been done in order to investigate the relationship between telomerase, telomere dysfunction and vulnerability to drug-induced apoptosis, particularly by cisplatin. Inhibition of telomerase activity or expression increased the susceptibility of glioblastoma cells,163 and of drug-resistant and drugsensitive promyelocytic leukemia and breast cancer cells164 to cisplatin-induced apoptosis. Results were controversial for melanoma cells, since neither an increase in sensibility for cisplatin was detected when telomerase was inhibited30 nor a precise relationship was found between telomerase activity and cellular sensitivity to cisplatin.137,165 Moreover, telomerase activity of cisplatin-treated cells has also been shown to depend on the period of treatment of prostate cell lines.166 Other studies revealed that the high level of hTERT expression and/or telomerase activity are related to chemosensitivity to cisplatin in ovarian carcinoma,167 and in oesophageal cancer cell lines,168 suggesting that telomerase could be a marker of prognosis for cisplatin treatment. A study of cisplatin treatment of telomere dysfunctional and telomere functional mice cells suggested that telomere dysfunction does not govern chemosensitivity to cisplatin.169 Based on all available data one may conclude that telomerase may play a role in tumour cell proliferation by modulating the cellular response to chemotherapy, particularly for drugs that affect DNA integrity.

7.9 Conclusion In vitro experiments have shown that telomerase and telomeres are good targets for cisplatin and some of its derivatives. The various results from cellular studies have been explained by differential suppression of hTERT expression and/or inhibition of telomerase activity that could lead to telomere shortening. However, this inhibition seems to depend on the cell lines, the doses and the experimental conditions. An unambiguous conclusion on the effects of cisplatin treatments may be obtained by comparing different cell lines under strict identical experimental conditions. Final conclusions must therefore await further studies. There is one way that has not yet been investigated extensively, but that appears to be of importance due to the increasing role of telomeres in cell viability; it concerns the possible platination of telomeric DNA. This event could lead to the wellknown cellular responses to cisplatin treatment leading to apoptosis or senescence.119 This hypothesis is supported by the fact that telomeric DNA is more accessible than genomic DNA94 and that telomere dysfunction induced by noncovalent quadruplex DNA binders is favoured in cancer cells over normal cells.33 Platination of telomeric

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DNA could lock the 3′- G-overhang in a G-quadruplex structure that is no longer recognized by telomerase; this could in turn inhibit replication of telomeres or uncap telomeres by impeding the binding of telomeric proteins. The latter approach is at present under investigation in our laboratory. Targeting telomere mechanisms could be of growing interest for future successful therapeutic anticancer strategies, since telomeres rather than telomerase may be the universal target for cancer therapy.

Acknowledgements We wish to thank Dr M.P. Teulade-Fichou, Dr D. Monchaud, H. Bertrand and Dr E. Segal-Bendirdjian for helpful discussions, and the Association pour la Recherche contre le Cancer (grants 3482 and 4835) for financial help.

Abbreviations TRF1 TRF2 POT1 TIN2 TPP1 Rap1 hTR hTERT 3′-G-overhang Pt-MPQ

Telomere repeat factor 1 Telomere repeat factor 2 Protection of telomere 1 TRF1 interacting nuclear protein 2 Tripetidylpeptidase 1 Repressor/activator protein 1 Human telomerase RNA component. Human telomerase reverse transcritase Single strand of the 3′ extremity of telomere that consists of G-rich repeated sequences Platinum–quinacridine complex

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157. Burger, A.M.; Double, J.A.; Newell, D.R.; Inhibition of telomerase activity by cisplatin in human testicular cancer cells; Eur. J. Cancer, 1997, 33(4), 638–644. 158. Raymond, E.; Sun, D.; Izbicka, E.; Mangold, G.; Silvas, E.; Windle, B. et al.; A human breast cancer model for the study of telomerase inhibitors based on a new biotinylatedprimer extension assay; Br. J. Cancer, 1999, 80(9), 1332–1341. 159. Akiyama, M.; Horiguchi-Yamada, J.; Saito, S.; Hoshi, Y.; Yamada, O.; Mizoguchi, H. et al.; Cytostatic concentrations of anticancer agents do not affect telomerase activity of leukaemic cells in vitro; Eur. J. Cancer, 1999, 35(2), 309–315. 160. Akeshima, R.; Kigawa, J.; Takahashi, M.; Oishi, T.; Kanamori, Y.; Itamochi, H. et al.; Telomerase activity and p53-dependent apoptosis in ovarian cancer cells; Br. J. Cancer, 2001, 84(11), 1551–1555. 161. Ku, W.C.; Cheng, A.J.; Wang, T.C.; Inhibition of telomerase activity by PKC inhibitors in human nasopharyngeal cancer cells in culture; Biochem. Biophys. Res. Commun., 1997, 241(3), 730–736. 162. Spiropoulou, T.; Ferekidou, L.; Angelopoulou, K.; Stathopoulou, A.; Talieri, M.; Lianidou, E.S.; Effect of antineoplastic agents on the expression of human telomerase reverse transcriptase beta plus transcript in MCF-7 cells; Clin. Biochem., 2004, 37(4), 299–304. 163. Kondo, Y.; Kondo, S.; Tanaka, Y.; Haqqi, T.; Barna, B.P.; Cowell, J.K.; Inhibition of telomerase increases the susceptibility of human malignant glioblastoma cells to cisplatininduced apoptosis; Oncogene, 1998, 16(17), 2243–2248. 164. Ward, R.J.; Autexier, C.; Pharmacological telomerase inhibition can sensitize drugresistant and drug-sensitive cells to chemotherapeutic treatment; Mol. Pharmacol., 2005, 68(3), 779–786. 165. Folini, M.; De Marco, C.; Orlandi, L.; Daidone, M.G.; Zaffaroni, N.; Attenuation of telomerase activity does not increase sensitivity of human melanoma cells to anticancer agents; Eur. J. Cancer, 2000, 36(16), 2137–2145. 166. Chen, Z.; Koeneman, K.S.; Corey, D.R.; Consequences of telomerase inhibition and combination treatments for the proliferation of cancer cells; Cancer Res. 2003, 63(18), 5917–5925. 167. Buttitta, F.; Pellegrini, C.; Marchetti, A.; Gadducci, A.; Cosio, S.; Felicioni, L. et al.; Human telomerase reverse transcriptase mRNA expression assessed by real-time reverse transcription polymerase chain reaction predicts chemosensitivity in patients with ovarian carcinoma; J. Clin. Oncol., 2003, 21(7), 1320–1325. 168. Asai, A.; Kiyozuka, Y.; Yoshida, R.; Fujii, T.; Hioki, K.; Tsubura, A.; Telomere length, telomerase activity and telomerase RNA expression in human esophageal cancer cells: correlation with cell proliferation, differentiation and chemosensitivity to anticancer drugs; Anticancer Res., 1998, 18(3A), 1465–1472. 169. Lee, K.H.; Rudolph, K.L.; Ju, Y.J.; Greenberg, R.A.; Cannizzaro, L.; Chin, L. et al.; Telomere dysfunction alters the chemotherapeutic profile of transformed cells; Proc. Natl. Acad. Sci. USA, 2001, 98(6), 3381–3386.

8 Towards Photodynamic Therapy of Cancer with Platinum Group Metal Polyazine Complexes David F. Zigler and Karen J. Brewer

8.1 Introduction This chapter focuses on the current state of the art of platinum group polyazine chromophores, their photochemistry with DNA and phototoxicity toward intact cells. It is intended to provide the basic principles and concepts for a researcher entering the field of inorganic photodynamic therapy (PDT). Early on the reader will be introduced to the basic principles of electronic excited states and PDT. In keeping with the theme of this book, special focus will be given to Ru(II), Os(II) and Rh(III) polyazine photochemistry with DNA. Comprehensive reviews have been published on the photochemistry of monometallic polyazine complexes with DNA,1,2 nonplatinum pharmaceuticals,3,4 photochemical action of metallointercalators and metalloinsertors of DNA,5 copper complex photochemistry with DNA,6 photoactive metal complexes in medicine7 and bioinorganic photochemistry.8 Many reviews have been published assessing the mechanism of PDT action at the tissue, cellular and molecular levels.9–11 This chapter will serve as an addendum to all of these reviews and include discussion of multimetallic complexes with highlights of the most recent work in this rapidly expanding field. Multimetallic and supramolecular systems are particularly promising as PDT agents due to the ease of tuning properties and reactivity by structural modification. The chapter will be punctuated by cell studies with inorganic polyazine chromophores, offering a glimpse into the Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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future of transition metal polyazine complexes as PDT agents. Polyazine metal complexes have properties that make them a promising class of future PDT agents, which will be highlighted herein. Photodynamic therapy is a modern treatment that uses light and a chromophoric molecule (PDT agent) to locally photosensitize tissues and to selectively kill targeted cells. PDT has potential applications in the treatment of localized tumours and intravascular diseases. Following administration of the PDT agent, the lesion of interest is irradiated with visible or near-infrared (NIR) light. Complicated photochemical reactions ensue, inducing apoptosis and/or necrosis, hopefully resulting in ablation of the lesion. PDT holds particular promise in the treatment of cancer as it tends to have fewer systemic toxic effects commonly associated with traditional radiation and chemotherapies. 8.1.1 Cancer Cancer is a disease caused by uncontrolled growth of abnormal cells, impacting the functioning of bodily tissues and organs. A mass of cancer cells, also known as a tumour or lesion, has many stages of growth. Initial mass formation is typified by rapid growth concomitant with poor vascularization within the lesion. Continued rapid growth results in the centre of the mass becoming hypoxic and undergoing necrosis. Abnormal cells may break off of the primary tumour and migrate throughout the body in a process called metastasis. Current cancer treatments are relatively crude, having severe side effects. These include surgery (invasive, risk of infection, complications), chemotherapy (poor cytospecificity/ selectivity, systemic toxicity effects) and radiation therapy (radiation sickness, increased risk of secondary tumours). Ideal treatments of cancer could be used in the ablation of lesions without the side effects which greatly impact the quality of life of the patient. 8.1.2 A Brief History of PDT and Current Clinical Treatments Photodynamic therapy has been expanding as a treatment of a wide range of localized maladies. Photodynamic action was first described in 1900 by Oscar Raab.12 In 1905 von Tappeiner and Jesionek reported the first successful clinical use of PDT to treat basal cell carcinoma with a topical application of eosin red and light.13 PDT studies were largely abandoned until the late 1970s, when animal trials using hematoporphyrin derivatives gave promising results in the treatment of animal cancers. Approval of the hematoporphyrin derivative, Photofin®, for use in PDT did not occur in the US until 1995 (Figure 8.1).14 Within the US, 5-aminolevulinic acid (Levulan®) is the only other approved chemical for use in PDT, gaining approval in 2000. Levulan® acts as a prodrug, enhancing porphyrin anabolism. The European Union, Norway and Iceland have approved Foscan®, a chlorin (chlorophyll derivative), for the treatment of head and neck cancers. At the time of writing there were three chromophores in current clinical trials: metoxifin lutetium (Lutex), tin ethyl

Introduction Me

R

Me Na O

Me Me

O H

N H N

237

N H N

N

N SnII N N

Me

O O

O NaO2C

n

hematoporphyrin derivative n = 1-4, Photofrin®

tin ethyl etiopurpurin Puryltintm OH

NH N

N

N

LuIII

N

HN

N

OH

tetraphenylchlorin Foscan® O

O

N

O

O

O

OMe

N

O

O

O

OMe

metoxifin lutetium Lutex

NH3+Cl-

OH

5-aminolevulinic acid Levulan® Figure 8.1 Chemical structures of compounds in clinical use and clinical trial for PDT in the United States15

etiopurpurin (Puryltin™) , and mono-L-aspartyl chlorine, with Lutex and Puryltin™ being the only inorganic centred chromophores in clinical trial.15 8.1.3 Characteristics of PDT Agents Ideal PDT agents absorb light strongly in the red or near-infrared (NIR) region of the spectrum, accumulate within the target tissue, efficiently perform the intended

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Towards Photodynamic Therapy of Cancer

action, have low systemic toxicity and are eliminated rapidly from the body. The red to NIR region of the spectrum (650–850 nm) is the light that has the greatest depth of penetration in human tissue and is known as the ‘therapeutic window’. Potent light absorbers with moderately long-lived and reactive electronic excited states are of particular interest. Chromophores with long-lived electronic excited states have a high probability of approach to a substrate during the excited state lifetime. Ironically, excited-state energy and lifetime often are inversely related, with longer wavelength light absorbers having short-lived excited states.16 Some current PDT agents show some cytospecificity, accumulating in tumour cells and enhancing the photosensitizer’s PDT function.14 Uptake of the PDT agent is often complicated and is impacted by the size of the chromophore, its charge and its lipophilicity. Chromophore size, charge and lipophilicity are important characteristics to consider in the design of PDT agents for elimination from the body. PDT agents that have hydrophobic architectures are retained for long periods within the body, leading to prolonged patient photosensitivity.15 Inorganic Complexes as PDT Agents Current PDT treatments employ organic chromophores as photosensitizers, but inorganic chromophores show great promise as PDT agents.1,8 The first approved PDT agent was a hematoporphyrin derivative, Photofrin® (Figure 8.1). Other photosensitizers have been approved, all of which are based on organic constructs.14 PDT has not gained widespread clinical use, owing in part to the newness of it as a therapy. In addition, the stigma of heavy-metal-containing compounds has led to limited use of these promising species as PDT agents. Two metal-containing photosensitizers are in clinical trials, metoxifin lutetium and tin ethyl etiopurpurin. Transition-metal-centred complexes with porphyrin, phthalocyanine, naphthalocyanine or polypyridine-type ligands have interesting ES properties, making them good targets for future therapies. This chapter will focus on inorganic systems with excited state properties and bioactivity that makes them of interest as future PDT agents.

8.1.4 DNA and PDT Action Deoxyribonucleic acid (DNA) is a natural supramolecule with complex secondary and tertiary structure whose integrity is central to the functioning of a cell (Figure 8.2). The importance of DNA to cellular function has directed research focus on transition metal complexes as potential PDT agents based on their ES reactions with DNA.2 Several structures of DNA have been reported with B-form DNA being the most prominent. A single turn of the B-form double helix is ca. 12 nm in length and averages 10.4 p-stacked base pairs.17 Two hydrophobic grooves run along the surface of the DNA molecule. The major groove is broad and shallow, and the minor groove is narrow and deep, both lined by the sugar phosphate backbone, giving DNA its polyanionic character. These structural features can be used in the rational design of DNA photomodification agents.

Study of Photophysics: Toward PDT 239

A H H N

T N

5' HO

N

O

O O P O O

3'

HN N O

O O P OO-

N N

O

N

O O P OO

N

H O

OO P O O

3'

N

N

O

-

O

H H

N

O

O

N

N H

NH

major groove

OH

5' C

minor groove

G

Figure 8.2 (Plate 10) Complimentary pairs of nucleotides that make up double helix DNA (Inset, A = adenine, T = thymine, G = guanine, C = cytosine, DNA = deoxyribonucleic acid). Atoms are carbon (grey), hydrogen (white), nitrogen (blue), oxygen (red) and phosphorous (pink) (See colour plate section)

8.2 Study of Photophysics: Toward PDT The discussion of transition metal polyazine photophysics requires explanation of general photophysical processes. Absorption of a photon by a chromophore causes an electronic transition and populates the chromophore’s electronic excited state. There are several internal pathways by which the excited chromophore might relax. The decay processes of electronically excited transition metal polyazine complexes have been the subject of study for decades, and continue to be a strong focus in research. Thorough studies of transition metal polyazine chromophores have led to many applications of the complexes in photochemistry. 8.2.1 Electronic Excitation Absorption of a photon of light by a chromophore in its electronic ground state (GS) is coincident with an electronic transition, populating an electronic excited state (ES) of the chromophore. The electronic transition promotes an electron from a low energy occupied orbital to an unoccupied orbital of higher energy (Figure 8.3). The lowest energy transition promotes an electron from the highest occupied molecular orbital (HOMO) to the lowest unoccupied molecular orbital (LUMO). The orbitals involved in the transition must be electronically coupled to allow electronic transition. The probability of an electronic transition is governed by selection rules. The symmetry selection rule for intensity of an electronic transition is expressed in Equation (8.1):

240

Towards Photodynamic Therapy of Cancer high energy electron LUMO hν

E

low energy hole HOMO 1

GS

1

ES

Figure 8.3 The change in electronic configuration following excitation by light of a singlet ground state chromophore (1GS) to a singlet electronic excited state (1ES) (HOMO = highest occupied molecular orbital, LUMO = lowest unoccupied molecular orbital) ES GS f ∝ ∫ ψ e1 µ ψ e1 dv

(8.1)

where f, the oscillator strength, is proportional to the integral of the product of the GS and ES wavefunctions (y)and m, the electric dipole moment operator. The spin selection rule states that for an electronic transition to be allowed, the spin multiplicity of the ground state and excited state must be the same. Electronic transitions which are allowed by donor–acceptor orbital electronic coupling, symmetry and electron spin are the most intense, with extinction coefficients, e, of ≥103 M−1 cm−1. Transitions which are allowed by spin, but formally forbidden by symmetry have e between 100 and 102 M−1 cm−1. Truly spin-forbidden transitions are not observed. Relaxation of the selection rules can give rise to observation of transitions that are formally forbidden. In the presence of a heavy atom, spin–orbit coupling relaxes the spin-selection rule, giving rise to absorptions that are formally spin forbidden displaying e as high as 103 M−1 cm−1. Electronic transitions often are accompanied by vibronic excitation. The conversion between electronic energy surfaces, moving from the 1GS to the 1ES, is dictated by the Franck–Condon principle (Figure 8.4). Electronic transitions following absorption of a photon, hn, are rapid, populating the 1ES in a nuclear configuration consistent with the 1GS. The 1ES is populated in a hot vibronic state (nn, where n is an integer >0). Vibronic relaxation to the lowest vibronic states is rapid with a rate constant kvib > 1012 s−1. Depopulation of the 1ES to the 1GS occurs at varying rates, with rate constants of the decay processes greatly depending on the nature of the 1 ES. 8.2.2 Unimolecular Electronic Excited State Decay The quantum efficiency of ES processes is an important description of the relative kinetics of ES decay. Decay from the ES by a specific pathway, x, is quantified by the quantum yield, Fx. The quantum yield is the probability of a molecule in its excited state decaying by a specific pathway. Quantum yield is defined as the ratio of the observed first-order rate constant of the process of interest, kx, divided by the sum of the rate constants of all pathways depopulating that state, Σk, Equation 8.2:

Study of Photophysics: Toward PDT 241

kvib > 1012s–1

ν6 ν5

ν7

1

ES

ν4 ν3

ν2 ν1 ν0

E

hν

kr = 103–1010s–1

hν' ν6

ν7

1

GS

ν5

ν4 ν3 ν2 ν1 ν0

reGS

reES

r Figure 8.4 Morse potential energy surface diagram for electronic (1GS, 1ES) and vibrational (νn) states of a chromophore. Included are typical first-order rate constants. kvib = vibronic relaxation rate constant, kr = rate constant of emission of light with average energy hν′, r = internuclear distance, reGS = GS equilibrium internuclear distance, reES = ES equilibrium internuclear distance.

Φ x = Φ ES

kx ∑k

(8.2)

where FES is the quantum efficiency of population of the reactive ES. An important descriptor of the electronic excited state is its inherent lifetime, t0, simply defined as (Σk)−1. Conversion between electronic states of systems is typically represented with Jablonski or state diagrams. These diagrams are simplified versions of the Morse potential energy surfaces, as shown in Figure 8.4, where each electronic state is represented as a line instead of a surface. Decay processes of electronic excited states, ES, may occur by multiple pathways (Figure 8.5). Radiative processes are typically represented as straight arrows ( ) and nonradiative processes are presented as wavy arrows ( ). Internal conversion, kic, is nonradiative relaxation without change in electron spin, while intersystem crossing, kisc, is accompanied by a change in spin. A molecular orbital depiction of the processes is shown in Figure 8.6. Fluorescence, kf, is defined as emission of a photon and decay to the ground state without change in electronic spin. Radiative decay with a change in spin state is defined as phosphorescence, kp. It should be noted that several examples in the literature incorrectly define fluorescence or phosphorescence by the lifetime of the

242

Towards Photodynamic Therapy of Cancer

1

ES

k isc 3

kf

E hν

ES

kp k ic k nr

1

GS

Figure 8.5 Jablonski diagram illustrating relative energies of states. Each bar ( ) represents the potential energy surface of an electronic state with lines above the state meant to reflect excited vibronic states. ES are excited states and GS is the ground state. Arrows represent electronic excited state decay via fluorescence (kf), internal conversion (kic), intersystem crossing (kisc), phosophorescence (kp) and nonradiative (knr) pathways

kisc

hν

E 1

GS

1

ES

3

ES

Figure 8.6 Orbital representation of intersystem crossing (kisc) from singlet excited state (1ES) to triplet excited state (3ES)

emitting ES. Nonradiative relaxation of an ES to the ground state is sometimes expressed by knr. 8.2.3 Electronic Excited States and Unimolecular Decay of Transition Metal Polyazine Transition metal polyazine complexes are potent light absorbers in the ultraviolet (UV) and visible region of the spectrum (Figure 8.7.)18 All polyazine complexes absorb UV light strongly due to intense internal ligand (IL) transitions of the aromatic ligands (Figure 8.8). IL transitions are called ligand centered (LC) in some literature. Ruthenium(II) and osmium(II) polyazine complexes absorb strongly in the visible region and have low-lying ES that are typically emissive, long-lived and reactive.19 Mononuclear Rh(III) polyazine complexes absorb strongly in the UV-A region due to IL transitions. The ES of rhodium(III) and dirhodium(II) polyazine complexes have reactivity that is strongly coupled to the type of polyazine ligands

Study of Photophysics: Toward PDT 243

np ns

t1u

a1g,t1u

σ*M

a1g

eg

σ*M

t1g,t2g, t1u,t2u π*

LF

(n-1)d

t2g,eg

t2g

IL

t1u,t2u,t1g,t2g a1g,t1u,eg

Metal Orbitals

L

t1u,t2u,t1g,t2g

π*

MLCT

πM

t1u,t2u,t1g,t2g

πL a1g,t1u,eg

σL

Molecular Orbitals

π

σ

Ligand Orbitals

Figure 8.7 Molecular orbital diagram of an octahedral metal complex, depicting common electronic transitions associated with transition metal (M) complexes with polyazine ligands (L) (IL = internal ligand, MLCT = metal to ligand charge transfer, LF = ligand field)

employed. Combining chromophores into mixed-metal supramolecules gives complexes with photophysical properties that are unique compared to the mononuclear analogues.20 Tuning the ES properties of the transition metal polyazine complexes allows for tuning of their photochemical properties. Table 8.1 summarizes the photophysical properties of transition metal polyazine complexes used in the study of photochemical degradation of DNA or light activated cell studies. Ruthenium(II) and Osmium(II) Polyazine Complexes Ru(II) and Os(II) complexes are interesting as photosensitizers as they possess reactive ES that are easily tuned by modification of their ligand coordination sphere.19 Mononuclear Ru(II) polyazine complexes typically have an absorption band in the 400–500 nm region due to the Ru(dp) → TL(p*) charge transfer transition (MLCT), where TL = terminal polyazine ligand (Figure 8.7). The electronic absorption spectrum of [Ru(bpy)3]2+ , bpy = 2,2′-bipyridine, is presented in Figure 8.9. The initially populated 1MLCT state undergoes intersystem crossing with unit efficiency (Fisc ≈ 1) to give the 3MLCT state (Figure 8.10).21 For tris(chelate) polyazine complexes of Ru(II), the 3MLCT state is long-lived at room temperature (t0 = 10– 1000 ns) and displays strong phosphorescence (Fp ≈ 0.10).19 The ES populated following metal-centred d → d excitation is the ligand field state (LF). Though higher in energy, the nonemissive 3LF state is thermally accessible at room temperature from the 3MLCT state. Thermal population of the 3LF state competes with phosphorescence or nonradiative decay pathways of the 3MLCT state as a decay pathway. Modification of the coordination environment of Ru(II) polyazine systems modulates the electronic properties of the chromophore. Bis(chelate) polyazine complexes of Ru(II) with a cis-Ru(TL)2 moiety tend to have absorption spectral

244

Towards Photodynamic Therapy of Cancer

Terminal Ligands N

N N

N

N

N

N

N

N

N

N

NH

N

N

N

NH

N

N

dppz

phi

R

N

N

NH

N

N

R

tpy

Ph2phen, R = H dsdp, R = SO3-

N

N

N

N

N

N

N

N

N

N

chrysi

NH

phzi

N

N

N

N

PHEHAT

ddz

NH

NH

N

R

phen, R = H Me4phen, R = Me

N

HAT

R

N

R

N

TAP

bpy R

N

DAP

Bridging Ligands

N

N

N

N

N

N

bpm

N

N

N

N

N

N

N

N

N

dpp

N

dpq

dpb

Figure 8.8 Polyazine ligands commonly used in the synthesis of transition metal complexes for the photomodification of DNA

features similar to the tris(chelate) systems. Emission quantum efficiencies from the 3 MLCT states of these complexes, however, are often orders of magnitude smaller. For example, the Fp of [Ru(bpy)3]2+ at room temperature is 0.06 versus Fp = 0.002 reported for [(bpy)2Ru(NH3)2]2+ at 157 K.19,22 This reduction in emission for [(bpy)2Ru(NH3)2]2+ is due to the more rapid interstate conversion between the

– 610 596 614 743 645

636 – – – 610 – – 692 690 698 – – – – –

345, 490 452 462 – 478g 466 437 452 552g 588g 682g 460 – – 480 474 486 277c 358 355 355 375

cis-[(bpy)2Ru(NH3)2](BF4)2 [Ru(bpy)3]Cl2 [Ru(phen)3]Cl2 [Ru(Ph2phen)3]Cl2 [Os(bpy)3](PF6)2 [(phen)Ru(HAT)2]Cl2

[Ru(TAP)3]Cl2

[(dppz)Ru(HAT)2]Cl2 [(bpy)2Ru(DAP)](PF6)2 [Ru(DAP)3](PF6)2 [Os(DAP)3](PF6)2 [(bpy)2Ru(ddz)]Cl2 [(phen)2Ru(ddz)]Cl2 Na4[Ru(dsdp)3] [(bpy)2Ru(dpp)]Cl2 [(phen)2Ru(dpp)]Cl2 [(Ph2phen)2Ru(dpp)]Cl2 [Rh(phen)3]Cl3 [(phen)2Rh(phi)]Cl3

cis-[(phen)2RhCl2]Cl cis-[(Me4phen)2RhCl2]Cl cis-[(5,6-Me2phen)2RhCl2]Cl

602

em (RT ) λmax (nm)

Complexb

abs λmax (nm)

723 704 710

– 698 727 763 – – – – – – 567 – – – –

1.090 – – – 0.180h 0.210h 3.600 – – – 4,100 –

–

0.052f 0.385 0.421 0.909 0.060 0.835

741f – – – 710 – –

t (RT) (µs)

em λmax (77 K )c (nm)

MLCT

3

LF LF 3 LF

3

3

MLCT, MLCT 3 MLCT 3 MLCT 3 MLCT 3 MLCT 3 MLCT 3 MLCT 3 MLCT 3 MLCT 3 IL 3 IL

3

3

3

LF MLCT 3 MLCT 3 MLCT 3 MLCT 3 MLCT

3

Lowest Energy ESd PB-M Type II-PC Type I & II-PC – – PB-L,Type I-PO Type II-PC PB-L,Type I-PO Type II-PC Type I-PO Type II-PC – – Type II-PC Type II-PC Type II-PC Type II-PC Type II-PC Type II-PC Type I-PC PB-M, Type I-PC PB-M PB-M PB-M

DNA Photochemistrye

30 66 66

72 77 77 77 76,81 76 76 75 75 75 26,73 58

50,67

22 2,56 2,56,74 2,56 24 72

Ref.

Table 8.1 Photophysical properties of some mononuclear transition metal polyazine complexes used in DNA photomodification schemesa

Study of Photophysics: Toward PDT 245

– – – – – – – – –

432 434 432 542 525 540 540 615 525

cis-[Rh2(m-O2CMe)2(dppz) (h1-O2CMe)(MeOH)](O2CMe) cis-[Rh2(m-O2CMe)2(dppz)2](O2CMe)2 cis-[Rh2(m-O2CMe)2 (dppz)(bpy)](O2CMe)2 [{(bpy)2Ru(dpp)}2 Ru(dpp)PtCl2](PF6)6 [{(bpy)2Ru(dpp)}2RhCl2](PF6)5 [{(bpy)2Os(dpp)}2RhCl2](PF6)5 [{(tpy)RuCl(dpp)}2RhCl2](PF6)3 [{(bpy)2Ru(bpm)}2RhCl2](PF6)5 [{(bpy)2Ru(dpp)}2IrCl2](PF6)5 – – – – –

–

– –

– – – – –

–

– –

–

–

554, 710i –

t (RT) (µs)

em λmax (77 K )c (nm)

3

MMCT MMCT 3 MMCT 3 MLCT 3 MLCT

3

MLCT

3

3

MMLCT MMLCT

3

MMLCT

3

IL, 3LF

3

Lowest Energy ESd

PC PC PC – –

Type II-PC

Type I & II-PC Type I & II-PC

Type I-PC, PB-M Type I & II-PC

DNA Photochemistrye

43 44 44 43 43

78

33 83

33

65

Ref.

b

Absorption maxima and room temperature emission maxima, reported in aqueous solutions, unless otherwise noted. bpy = 2,2′-bipyridine; phen = 1,10-phenanthroline; Ph2phen = 4,7-diphenyl-1,10-phenanthroline; dppz = dipyrido[3,2-a:2′,3′-c]phenazine; HAT = 1,4,5,8,9,12-hexaazatriphenylene; TAP = 1,4,5,8-tetraazaphenanthrene; DAP = 1,12-diazapyrylene; ddz = dibenzo[h,j]dipyrido[3,2-a:2′,3′-c]phenazine; dsdp = 4,7-di(3-sulphurylphenyl)-1,10-phenanthroline; dpp = 2,3-bis(2-pyridyl)pyrazine; phi = 9,10-phenanthrenequinone diimine; Me4phen = 3,4,7,8-tertramethyl-1,10-phenanthroline; 5,6-Me2phen = 5,6-dimethyl-1,10-phenanthroline, tpy = 2,2′:6′,2″-terpyridine; bpm = 2,2′-bipyrimidine c In 4:1 EtOH/MeOH d LF = ligand field, MLCT = metal-to-ligand charge transfer, IL = internal ligand, MMLCT = metal-metal to ligand charge transfer, MMCT = metal to metal charge transfer 5 PB-L = photobinding to ligand, PB-M = photobinding to metal, Type I = direct ES & DNA reaction following electron transfer, Type II = photosensitized oxygen mediated reactions, PC = photocleavage, PO = photooxidation d 157 K. g In room temperature acetonitrile h Multiexponential, reported as the average of three components i Emission is observed from both 3LC and 3LF states in 4 : 1 EtOH/MeOH glass

a

–

em (RT ) λmax (nm)

380

abs λmax (nm)

cis-[(phen)(dppz)RhCl2]Cl

Complexb

Table 8.1 Continued

246 Towards Photodynamic Therapy of Cancer

Study of Photophysics: Toward PDT 247 8

bpy(π→π*)

ε x10-4 (M-1cm-1)

7

N

6

N

5

N RuII N

N N

4 3 2

Ru(dπ)→bpy(π*) CT

1 0 200

300

400

500

600

Wavelength (nm)

Figure 8.9 Electronic absorption spectrum of tris(2,2′-bipyridine)ruthenium(II), [Ru(bpy)3]2+, in CH3CN at RT

1

IL

∆E (x103 cm-1)

30

kic

25 20

1

kisc

MLCT

3

LF

3

MLCT knr hν' kp hν

15 10

knr'

5 0

1

GS

Figure 8.10 Jablonski-type diagram of [Ru(bpy)3]2+. bpy = 2,2′-bipyridine, 1GS = singlet electronic ground state, 1MLCT = singlet metal to ligand charge transfer excited state, 3MLCT = triplet MLCT excited state, 1IL = singlet internal ligand excited state, 3LF = triplet ligand field excited state. Relative energies are adapted from Juris, Balzani, Barigletti, Campagna and Belser.19

3

MLCT state and lower-lying 3LF state (Figure 8.11). Excitation directly into higher LF states has been linked to photoaquation of complexes with the cis-RuII(TL)2 moiety, where complexes such as [(bpy)2RuCl2] photochemically generate [(bpy)2RuCl(OH2)]+ and [(bpy)2Ru(OH2)2]2+.23 Tris(chelate) polyazine complexes of Os(II) generally absorb and emit visible light, but at longer wavelengths and with decreased emission efficiency than the Ru(II)-centred analogs.24,25 A shorter 3MLCT state lifetime (and smaller Fp) is a result of several contributing factors. The energy gap law states that as the difference in energy of two electronic states decreases, the vibronic coupling of the states increases, enhancing knr and leading to more efficient ES deactivation.24 Another factor to consider is the large spin–orbit coupling of Os(II) versus Ru(II). A larger spin–orbit coupling constant relaxes the spin selection rule, increasing the intensity of spin forbidden transitions. This gives rise to a low energy ‘tail’ of the visible region

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Towards Photodynamic Therapy of Cancer 1

∆E (x103 cm-1)

30 25 20

1 1

IL LF

kisc'

3

kisc

MLCT

3

3

MLCT knr hν' k

15

nr'

hν

10

kp

LF2 LF1

knr''

krxn

krxn'

5 1

0

GS

Figure 8.11 Jablonski-type diagram of [(bpy)2Ru(NH3)2]2+. bpy = 2,2′-bipyridine, 1GS = singlet electronic ground state, 1MLCT = singlet metal to ligand charge transfer excited state, 3 MLCT = triplet MLCT excited state, 1IL = singlet internal ligand excited state, 3LF = triplet ligand field excited state. Relative energies are adapted from Singh and Turro22

1

IL

∆E (x103 cm-1)

30 kic

25 20

1

MLCT

15

3

LF

kisc 3

hν' hν

10

hν'' kp

MLCT

knr

5 0

1

GS

Figure 8.12 Jablonski-type diagram of [Os(bpy)3]2+. bpy = 2,2′-bipyridine, 1GS = singlet electronic ground state, 1MLCT = singlet metal to ligand charge transfer excited state, 3MLCT = triplet MLCT excited state, 1IL = singlet internal ligand excited state. Relative energies are adapted from Kober, Caspar, Lumpkin and Meyer24

electronic absorption spectra of Os(II) polyazine complexes. The low energy tail is due to direct population of the 3MLCT state (Figure 8.12).24 The larger spin–orbit coupling constant also enhances decay of the 3MLCT state to the 1GS. Unlike the Ru(II) counterparts, Os(II) polyazine 3LF states are not thermally accessible from the 3MLCT state. Therefore, excitation of the Os(II) chromophore with visible light generally does not result in ligand substitution. Chromophores based on Ru(II) and Os(II) polyazine complexes have found great utility in the study of DNA photomodification. Rhodium Polyazine Complexes Rhodium polyazine complexes have distinct, reactive electronic excited states that make them attractive as PDT agents. As with other transition metal polyazine complexes discussed in this chapter, the ligand environment around the central rhodium has major impact on the ES properties. Mononuclear tris(chelate) Rh(III) polyazine complexes are potent light absorbers in the UV region of the spectrum due to strong

Study of Photophysics: Toward PDT 249 1

35

IL

kisc

∆E (x103 cm-1)

30

3

IL

25 hν

20

kp

15

knr

10 5 1

0

GS

Figure 8.13 Jablonski-type diagram of [Rh(phen)3]3+. phen = 1,10-phenathroline, 1GS = singlet electronic ground state, 1IL = singlet internal ligand excited state. Relative energies are adapted from Crosby and Elfring26

1

kisc'

IL LF

30

∆E (x103 cm-1)

1

25

3

IL

kisc

20 15

hν

0

hν'

knr

5 1

LF2

3

LF1

kic kp

10

3

krxn

GS

Figure 8.14 Jablonski-type diagram of [(phen)2RhCl2] +. phen = 1,10-phenathroline, 1GS = singlet electronic ground state, 1IL = singlet internal ligand excited state, 3LF = triplet ligand field excited state. Relative energies are adapted from Demas and Crosby27

IL transitions. These Rh(III) complexes lack the visible light absorption bands as their MLCT transitions occur at much higher energy than analogous Ru(II) and Os(II) systems (Figure 8.13).26 Coincidently, the complexes emit from a 3IL state. Bis(chelate) polyazine complexes of Rh(III) with a cis-RhIIIX2 moiety, where X is a halide, also absorb only in the UV and are emissive in low temperature glasses.27 A weak absorption band observed between 350–400 nm arises from LF transitions for the [(TL)2RhX2]+ systems. The emission from these complexes, when excited into the IL or LF bands, is broad and significantly Stokes shifted relative the lowest absorption band (∼14 000 cm−1).28 The initially populated 1LF state intersystem crosses to an upper 3LF2 state (Figure 8.14). The 3LF2 state decays by internal conversion to the emissive 3LF1 state. At room temperature, photoaquation (krxn) competes with nonradiative decay to give the mono- and disubstituted solvato complexes, [[(TL)2RhX(S)]2+ and [(TL)2Rh(S)2]3+ (S = solvent).23,29 Morrison and coworkers have exploited this decay pathway to photobind to DNA, mimicking the

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Towards Photodynamic Therapy of Cancer

∆E (x103 cm-1)

30 25 20

1

IL

kisc

MMLCT

kisc'

3

MM-MM

kisc"

3

1 1

15

3

MM-MM

10

hν knr

5 0

IL MMLCT krxn

1

krxn'

GS

Figure 8.15 Jablonski-type diagram of cis-[Rh2(m-O2CCH3)2(dppz)2]2+. dppz = dipyrido[3,2a:2′,3′-c]phenazine, 1GS = singlet electronic ground state, 1IL = singlet internal ligand excited state, 1MMLCT = singlet metal–metal to ligand charge transfer state, 1MM-MM = Rh-Rh(dp*dΣ*) state. Relative energies are adapted from Angeles-Boza, Bradley, Fu, Wicke, Bacsa, Dunbar, Turro33,82

DNA thermal binding properties of the well known complex cis-[Pt(NH3)2Cl2] (cisplatin).30,31 Dirhodium(II) complexes possess unique ES compared to the mononuclear Rh(III) complexes due to the strong electronic coupling of the two metal centres. Complexes of the form [Rh2(m-O2CCH3)4] absorb weakly in the near UV region owing to a low lying Rh-Rh(dp*) → Rh-Rh(ds*) (MM-MM) transitions (Figure 8.15).32 Inclusion of ligands with stabilized p*-acceptor orbitals, e.g. dibenzo[h,j]dipyrido[3,2-a:2′,3′-c]phenazine (dppz), gives complexes with transitions at ∼430 nm attributed to IL transitions and Rh-Rh(dp*) → dppz(p*) charge transfer transitions (MMLCT).33 The 3MMLCT state is thought to be a long-lived state with an oxidized dirhodium core. These complexes display DNA photooxidation reactions, consistent with reaction of the 3MMLCT state. Excited States of Polyazine Bridged Ru–Rh Complexes Polyazine bridged, mixed-metal supramolecules have been developed that incorporate multiple metal centres and have more complicated photophysics than the mononuclear analogues.34 The mixed-metal complexes often retain some ES properties of the subunits used to assemble the supramolecule. Metal centres with strong electronic coupling, e.g. di-Rh(II) complexes above, have electronic transitions that involve orbitals delocalized over the metal centres. At the other extreme, metal centres that are covalently linked, but without strong electronic coupling, behave like the mononuclear analogues. Ru(II)–Rh(III) dyads linked via an alkyl tether have electronic properties consistent with each subunit, indicating weak electronic interaction between coupled chromophores.35–39 The polyazine bridged complexes have intercomponent interactions, giving rise to states that are unique to the supramolecular complex without full delocalization of orbitals. Mixed-metal supramolecules that have weakly to moderately coupled metal centres exhibit unique ES with interesting ES dynamics, giving rise to useful pho-

Study of Photophysics: Toward PDT 251

1

∆E (x103 cm-1)

30

IL kic

25 20 1MLCT 15

kisc 3

hν

10

MLCT kp

ket knr

5 0

3

krxn

MMCT

knr'

1

GS

Figure 8.16 Jablonski-type diagram of [{(bpy)2Ru(dpp)}2RhCl2]5+. bpy = 2,2′-bipyridine, dpp = 2,3-bis(2-pyridyl)pyrazine, 1GS = singlet electronic ground state, 1MLCT = singlet metal to ligand charge transfer, 3MLCT = triplet MLCT, 1IL = singlet internal ligand excited state. Relative energies adapted from Molnar, Jensen, Vogler, Jones, Laverman, Bridgewater, Richter and Brewer.41

tophysical properties. Metals bridged by 2,3-bis(2-pyridyl)pyrazine (dpp) electronically couple through the aromatic pyrazine ring.40 Mononuclear Ru(II) complexes with dpp have a strong, visible MLCT transition and a LUMO localized on the abs pyrazine ring of the dpp ligand (e.g. [(bpy)2Ru(dpp)]2+, λmax = 480 nm in water). Bridging the complex to another metal centre stabilizes the dpp(p*) LUMO, shifting abs the MLCT transition to lower energy (e.g. [(bpy)2Ru(dpp)Ru(bpy)2]4+, λmax = 526 nm ). II III Trimetallic complexes with Ru (dpp) subunits bridged to a cis-Rh Cl2 centre such as [{(bpy)2Ru(dpp)}2RhCl2]5+ have interesting properties (Figure 8.16).41 The HOMO of these trimetallic complexes is localized on the Ru(dp) orbitals like other Ru(II) polyazine complexes, but the LUMO is Rh(ds *) in nature. Direct Ru(dp) → Rh(ds*) (metal-to-metal) charge transfer (MMCT) excitation is forbidden by symmetry and the relatively weak HOMO–LUMO electronic coupling. Irradiation with visible light, however, populates the 1MLCT state. Intersystem crossing generates the 3 MLCT state, which is higher in energy than the 3MMCT state.42 Coupling of the dpp(p*) and Rh(ds *) acceptor orbitals facilitates intramolecular electron transfer to give the 3MMCT state. These trimetallic complexes have been shown to photocleave DNA.43,44 Substitution of the bridging ligand dpp for molecular bridges with lower energy acceptor orbitals decreases the photochemical reactivity, implicating the 3MMCT state.44 8.2.4 Bimolecular Excited State Interactions Ground state light absorbers (LA) and their electronic excited states (*LA) are interesting for study, but it is the interaction of *LA with substrate molecules that interests PDT researchers. Bimolecular interactions resulting in quenching of *LA following excitation (Equation 8.3) and competing with unimolecular radiative decay (Equation 8.4) and nonradiative decay (Equation 8.5) can be classified into basic groups: bimolecular deactivation, knr ′ (Equation 8.6); excited state electron transfer, ket, to (Equation 8.7) or from (Equation 8.8) *LA; excited state energy

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transfer, ken (Equation 8.9); and photochemical reaction, krxn (Equation 8.10); where Q = quencher, ED = electron donor, EA = electron acceptor. Ia LA + hν  → *LA

(8.3)

kr *LA  → LA + hν ′

(8.4)

knr *LA  → LA

(8.5)

knr′ *LA + Q  → LA + Q

(8.6)

ket *LA + ED  → LA − + ED+

(8.7)

ket′ *LA + EA  → LA + + EA −

(8.8)

ken *LA + Q  → LA + *Q

(8.9)

krxn *LA + reactant  → photoproducts

(8.10)

Current research efforts toward PDT agents study the photochemistry of an excited chromophore with DNA. Transition metal polyazine complexes have a rich history of photochemistry with DNA, focusing strong research efforts. These efforts have given rise to recent discoveries of previously unknown photochemistry. The field is expanding rapidly to accommodate increasing interest in DNA photochemical modification and inorganic-based PDT. In addition, information about fundamental photophysical properties is being uncovered. Excited State Electron Transfer Theory Reductive and oxidative quenching of *LA is driven by the relative redox potentials of the ED or EA and *LA. The *LA reduction E(*LAn/LAn−1) and *LA oxidation E (*LAn/LAn+1) potentials are estimated with the following equations: E(*LA n LA n−1 ) ≈ E(LA n LA n−1 ) − ∆E 0−0

(8.11)

E ( *LA n LA n+1 ) ≈ E ( LA n LA n+1 ) + ∆E 0−0

(8.12)

where E(LAn/LAn−1) and E(LAn/LAn+1) are the respective first reduction and oxidation potentials in V of the GS chromophore, and ∆E0−0 is the difference of energy between lowest vibronic states of the GS and ES in eV (see Figure 8.4). The ∆E0−0 em is commonly estimated from the λmax with a Stokes shift applied to account for the shift of the excited state energy surface relative to the ground state. The rate of ES quenching is governed by the driving force of electron transfer as described by Marcus.45 For a bimolecular reaction under pseudo-first order conditions:

[ ED]o or [ EA ]o >> [ ES]

Study of Photophysics: Toward PDT 253

the observed rate constant of electron transfer (kobs,et) is defined as: kobs,et = vNκ exp ( −∆G‡ RT )

(8.13)

where vN is the nuclear frequency factor and k is the electron transfer coefficient. The free energy of activation, ∆G‡, is expressed as:

∆G‡ =

λ ∆G0    1 + 4 λ 

2

(8.14)

where l is the reorganizational energy and DG0 is the change in standard free energy of redox reactants and products. Experimentally ket can be determined by Stern– Volmer kinetic analysis as described below. Type I Photooxidation Reactions in PDT Reductive quenching of an excited photosensitizer by DNA constitutes a Type I photooxidation reaction, as defined by Foote.46 For example, consider the mechanism of decay of the 3MLCT state of [(phen)Ru(HAT)2]2+ in the presence of DNA, phen = 1,10-phenanthroline, HAT = 1,4,5,8,9,12-hexaazatriphenylene. 2+ a → 1 * [( phen) Ru ( HAT )2 ] [( phen) Ru ( HAT )2 ]2 + + hv I 1

3

kisc 2+  → 3 * [( phen) Ru ( HAT )2 ]

(8.16)

knr 2+  → [( phen) Ru ( HAT )2 ]

(8.17)

kp 2+  → [( phen) Ru ( HAT )2 ] + hv ′

(8.18)

2+

* [( phen) Ru ( HAT )2 ] 3

2+

* [( phen) Ru ( HAT )2 ] 2+

* [( phen) Ru ( HAT )2 ]

(8.15)

2+ 2+ q * [( phen) Ru ( HAT )2 ] + DNA k → [( phen) Ru ( HAT )2 ] + DNA

(8.19)

2+ + et * [( phen) Ru ( HAT )2 ] + DNA k → [( phen) Ru ( HAT )2 ] + DNA +

(8.20)

Kinetically the quantum yield of the electron transfer reaction (Equation 8.20) is expressed as:

Φ et = Φ isc

ket [ DNA ] kp + knr + ( ket + kq ) [ DNA ]

(8.21)

where Fet is the quantum yield of electron transfer, Fisc is the efficiency of population of the 3MLCT state, kp and knr are the respective unimolecular phosphorescence and nonradiative decay rate constants, ket is the rate constant of electron transfer, kq is rate constant of bimolecular deactivation and [DNA] is the effective concentration of the DNA quencher in molarity of base pairs. Efficiency and rates of excited state reactions can be determined by probing the quantum yield of phosphorescence Fp:

Φ po = 1 + τ o (ket + kq ) [ DNA ] Φp

(8.22)

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Towards Photodynamic Therapy of Cancer

with Fp being the quantum yield of phosphorescence in the presence of quencher and Φ p0 is the phosphorescence yield in the absence of quencher. Stern–Volmer analysis consists of plotting Φ p0 Φ p as a function of [DNA] times the ES lifetime t0 in the absence of quencher, providing a linear relationship with a slope equal to kSV, where kSV = ket + kq. In the absence of measurable ES emission, ket may be determined by observing the changing in ES lifetime as measured by transient absorption spectroscopy because:

Φ p0 Φ p = τ τ 0

(8.23)

where t is the excited state lifetime in the presence of quencher. Transient spectroscopy can differentiate electron transfer quenching and other decay pathways probing the formation of the electron transfer product. Measuring the rate of formation of the product can also allow for the determination of ket. Excited State Energy Transfer Theory Two mechanisms of excited state energy transfer (Equation 8.9) have been described and involve either ES dipole resonance coupling or electron exchange between the *LA donor and quencher, Q (Equation 8.9). These two mechanisms have been described individually by Förster47 and Dexter.48 Förster described *LA quenching by excited state energy transfer through space by a Q.47 Resonant coupling of the chromophore and quencher excited state dipoles allow for radiationless energy transfer from the *LA donor to the acceptor Q. Förster energy transfer efficiency is governed by the spectral overlap of *LA emission and Q absorption, interchromophore distance and relative dipole orientation. Förster excited state energy transfer is the dominant pathway when *LA → LA and Q → *Q transitions are spin allowed. Förster excited state energy transfer is the governing principle of fluorescence resonance energy transfer (FRET) techniques used in confocal microscopy. Another mechanism of excited state energy transfer, described by Dexter, involves electron exchange between the *LA and Q.48 Electron exchange requires some electronic coupling of the *LA donor and the Q acceptor orbitals. Dexter transfer is not bound by the spin selection rule and is thought to be the dominant mechanism of *LA energy transfer when there is poor spectral overlap of *LA emission and Q absorption, e.g. photosensitization of 3O2 → 1O2. Study of excited state energy transfer quenching dynamics can be performed using Stern–Volmer kinetic analysis. Details of this analysis are outlined above for excited state electron transfer. In some instances, observation of the rise-time and/or decay of the *Q state can also be observed and aids in understanding this bimolecular interaction for a specific system. Type II Photooxidation Reactions in PDT Type II photooxidation reactions of biomolecules, as defined by Foote,46 involves photosensitization of molecular oxygen (3O2), generating singlet oxygen (1O2) which reacts with a biologically important substrate molecule. Photosensitization of 1O2

Photochemical Reactions of Metal Complexes with DNA

255

from 3O2 requires excited state energy transfer by the Dexter mechanism. The 3 MLCT state of [Ru(phen)3]2+, has been shown to produce 1O2 from 3O2 (Equation 8.28).49 a → [Ru ( phen)3 ]2 + + hv I 1

* [ Ru ( phen)3 ]

2+

3

3

3

* [ Ru ( phen)3 ]

1

* [ Ru ( phen)3 ]

2+

(8.24)

kisc 2+  → 3 * [ Ru ( phen)3 ]

2+

knr  →

(8.25)

[Ru ( phen)3 ]2 +

(8.26)

2+ 2+ p *[ Ru ( phen )3 ] k → [ Ru ( phen )3 ] + hv′

2+ ken * [ Ru ( phen)3 ] + 3 O2  →

[Ru ( phen)3 ]2 + +

(8.27) 1

O2

(8.28)

knr′ → 3 O2 O2 

(8.29)

kp′ O 2  → 3 O2 + hv ′′

(8.30)

krxn DNA + 1 O2  → DNA photoproduct

(8.31)

1

1

Using the steady-state approximation on the [1O2], the quantum efficiency of oxygen mediated photomodification of DNA (Frxn) can be described by: ken [ 3 O2 ] krxn [ DNA ]    Φ rxn = Φ isc   kp + knr + ken [ 3 O2 ]   knr′ + kp′ + krxn [ DNA ] 

(8.31)

where Fisc is the quantum yield of population of the 3MLCT state, ken is the rate constant for ES energy transfer to oxygen, 3O2, dissolved in solution and krxn is the rate constant for the reaction of 1O2 with DNA, where [DNA] is expressed as concentration of DNA base pairs. The rate constant ken is determined experimentally by measuring emission from the 3MLCT state of [Ru(phen)3]2+ in the absence of DNA, at different concentrations of 3O2. The rate constant krxn can be elucidated from DNA photoproduct analysis or by observing 1O2 emission decay.

8.3 Photochemical Reactions of Metal Complexes with DNA DNA plays important roles in the functioning of cells because it is central to protein expression (transcription) and cell replication. Damage to DNA manifests itself in many forms, some of which are detrimental to the integrity of the data stored on the DNA. Damage to DNA disrupts cell function by causing carcinogenesis, mutagenesis, apoptosis and cell death. There are several reported mechanisms of excited state photosensitizer reactions inducing DNA damage.2,5,50–53 Careful design of DNA photochemical experiments can yield surprising amounts of information about the mechanism of DNA/metal complex interaction as well as information about the

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Towards Photodynamic Therapy of Cancer

chromophore’s excited state dynamics. Characteristic changes to DNA are observed for photobinding of metal complexes, photooxidation of nucleobases and photocleavage of the sugar phosphate backbone. Knowledge of the decomposition pathways of DNA photolyzed with a metal complex helps to understand a possible mechanism of PDT action. 8.3.1 DNA Targets and Analysis Study of the photochemical reactions of metal complexes with DNA requires careful choice of the type of DNA used and method of analysis. Helpful descriptions of DNA structure and analysis can be found in the book by Calladine.17 Mono- and oligonucleotides are used as targets when chemical analysis techniques such as mass spectrometry (MS) and nuclear magnetic resonance (NMR) spectroscopy are used to study the photoproducts. Mononucleotides are available commercially and have found utility in experiments to understand binding of metal complexes to DNA and ES quenching studies. Oligonucleotides (oligos) are short strands of DNA (15–100 base pairs) that have known sequence and structural properties. Oligos are better models for nuclear DNA because they can be obtained as double stranded DNA. High-resolution poly(acrylamide) gel electrophoresis (HR-PAGE) is instrumental in understanding sequence specificity of photochemical reactions with oligonuclear DNA leading to irreversible changes. Circular plasmid DNA extracted from bacteria is obtained with a supercoiled tertiary structure that makes this type of DNA an excellent target for photomodification schemes.54,55 Changes to the topographical nature of supercoiled circular plasmid DNA are indicative of several decomposition pathways that can be analysed using agarose gel electrophoresis (Figure 8.17). Finally, genomic DNA is commercially available as a freeze dried extract from various tissue sources and is high molecular weight, with > 106 base pairs per molecule. Genomic DNA is used to understand DNA–metal interactions that require large volumes and/or high DNA concentrations for analysis using selective precipi-

λ Cir Lin 23.1 kbp9.4 kbp6.6 kbp4.4 kbp-

Form II

2.3 kbp2.0 kbp-

Form III Form I

Figure 8.17 Photograph of a typical 0.8% agarose gel illustrating the electrophoretic migration (moving from top to bottom) of three different forms of plasmid DNA. The lanes from left to right are as follows: l is a lambda DNA/HindIII digest molecular weight marker (kbp = 1,000 base-pairs), Cir is circular plasmid DNA containing native supercoiled (Form I) and open-circular (Form II) plasmid DNA, and Lin is linear plasmid DNA (Form III). Band assignments are based on Vinograd and Lebowitz55

Designing Transition Metal Polyazines for Photomodification

257

tation, changes to spectroscopy and viscosity measurements. Each form of DNA has specific advantages and disadvantages toward use in photochemical experiments and multiple types are required to get a more complete understanding of the metal complex–DNA photochemical mechanism.

8.4 Designing Transition Metal Polyazines for DNA Photomodification Photomodification of DNA by transition metal polyazine complexes is attractive because of the structural diversity of this class of complexes.18,19 Two major design considerations for chromophores applicable as DNA photomodification agents are: the impact of structural changes on the interaction of the chromophore with DNA, and the impact of chromophore structure on the efficiency and mechanism of their DNA photochemistry. Recent reports present photochemical mechanisms that help elaborate previously described photochemistry (Table 8.1) and have prompted cell studies to explore application in PDT. 8.4.1 Ground State Interactions of Transition Metal Polyazine Complexes and DNA The mechanism of DNA modification by photosensitizers is often multistep with the first step involving some form of metal complex–DNA association.2,50,51 Smaller, compact polyazine transition metal complexes tend to have an electrostatic interaction with DNA.56 Lipophilic substituents on the polyazine ligands give complexes that bind in the hydrophobic minor or major groove of the DNA double helix.57 Ligands with extended, planar aromatic substituents favour intercalation between DNA bases.5,51 More recently, metal complexes have been shown to insert between strands of the double helix, a behaviour that is previously not described.51 Binding of transition metal polyazine complexes to DNA is enantioselective, a reflection of the chiral nature of DNA and the often chiral nature of polyazine chelated metal complexes. Barton has pioneered exploiting the different binding modes as shapeselective probes of DNA and RNA structure.5,51,58,59 The strength of the DNA–metal complex interaction has a direct impact on the photosensitizer function as a DNA modification tool. Electrostatic interaction of cationic transition metal polyazine complexes with DNA can be described by the polyelectrolyte approximation of counterion condensation theory.60 DNA is polyanionic with a single negative charge per nucleotide in aqueous solution. Usually Na+ acts as the counterion to the DNA negative charge. Transition metal polyazine complexes studied for DNA photomodification can have large negative to large positive charges. Exchange efficiency of Na+ at the DNA surface by an incoming complex ion is a factor of the charge of the complex, the concentration of free metal complex and Na+ concentration (ionic strength). Complexes with large positive charges that are structurally compact have stronger electrostatic interactions with DNA. The prototypical transition metal polyazine complex [Ru(bpy)3]2+ exhibits ionic binding to DNA.56

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Groove binding of transition metal polyazine complexes to DNA is controlled by complex lipophilicity and molecular shape. Also known as surface binding, groove binding is association of a species with the hydrophobic pocket formed between the sugar-phosphate backbones of each strand. The complex [Ru(Me4phen)3]2+ is a wellknown groove binder.57 The methyl substituents on the phen ligand contribute to the complex’s overall lipophilicity, but provide steric bulk that prevents more intimate interactions with the DNA. Extended, planar polyazine ligands that can intercalate into the base stack of DNA have been shown to enhance metal complex–DNA electronic interactions prior to photolysis.5,51 Two ligands often included in a metal complex’s architecture that promote intercalation are dppz and 9,10-phenanthrenequinone diimine (phi). Complexes with bulky planar ligands have been reported to bind by an alternative mechanism. The complex [(phen)2Rh(chrysi)]3+ reported by Barton and coworkers inserts its a-chrysenequinone diimine (chrysi) ligand into the bases stack accompanied by displacement of a base pair. Photocleavage of DNA by metalloinsertor complexes was shown to be an effective method of identifying base mismatches in the DNA.61 It is important to note that the mode of DNA–metal complex interaction prior to photolysis is not always clear. The reader is referred to a review by Kirsch DeMesmeaker, Lecompte and Kelly for a historical account of ‘The controversial case of [Ru(phen)3]2+.2 Hexacoordinate transition metal complexes with two or three bidentate ligands tend to have strongly enantioselective binding interactions with DNA.59 DNA in its most common B form is a right-handed double helix (Figure 8.2), interacting more strongly with ∆ enantiomers of tris(chelate) metal complexes. Barton has shown that the complex ∆-[Ru(Ph2phen)3]2+ binds strongly in the major groove of the doublehelix, intercalating the base pairs. The other enantiomer Λ-[Ru(Ph2phen)3]2+ does not bind as strongly, owing to the steric repulsion of the bulky ancillary ligands with the sugar-phosphate backbone of DNA provided by the stereochemistry around the metal centre.

8.4.2 Mechanisms of DNA Photochemical Degradation Photochemical degradation of DNA has been pursued in recent years toward applications in PDT, and DNAase and RNAase biomimetics. The mechanisms of DNA degradation under photolytic or oxidative stress have been thoroughly reviewed, so minimal explanation of these pathways is given below.52,53,62 Photoinitiated binding of metal complexes has been studied, coined ‘photo-cisplatin’ agents. Photobinding through an ancillary ligand also leads to DNA covalent modification. Oxidation of the DNA bases by ES electron transfer reaction and hydrogen atoms abstraction are important to the photochemical function of some transition metal polyazine complexes. Photocleavage of DNA by the transition metal polyazine ES in the presence or absence of dissolved 3O2 reflects separate mechanisms of action. Many of the transition metal polyazine complexes studied for their photochemical interaction with DNA exhibit multiple ES reactions with DNA.

Designing Transition Metal Polyazines for Photomodification

259

Design Aspects: Toward Photobinding DNA Metal complexes can be designed to bind DNA after excitation. The variety of ES and their photochemistry provide a range of mechanisms for DNA photobinding. Photobinding DNA through a metal centre. Ruthenium(II) complexes of the form cis-[(TL)2RuX2]n+, where X = halide (n = 0) and X = H2O, NH3 or aliphatic amine (n = 2), have been investigated for their DNA thermal binding properties and more recently as DNA photobinding agents.3 The complexes cis-[(bpy)2RuCl2] and cis-[(bpy)2Ru(OH2)2]2+ readily covalently bind to DNA under physiological conditions. Photobinding to DNA of cis-[(bpy)2RuCl2] and cis-[(bpy)2Ru(OH2)2]2+ shows little enhancement over thermal binding pathways.63 Singh and Turro reported the DNA photobinding properties of cis-[(bpy)2Ru(NH3)2]2+, but have shown that the complex did not appreciably thermally bind DNA.22 The mononuclear complex absorbs strongly at 290 nm and 243 nm, consistent with IL transitions. abs MLCT transitions in the visible region occur at λmax = 490 nm . The complex ex undergoes photoaquation when irradiated with near-UV light (λmax = 350 nm or 400 nm, Faq = 0.024 ± 2 or 0.018 ± 2). Excitation with light > 450 nm, however, did not lead to observable chemical reaction. The authors indicate the photoaquation takes place from a 3LF state that is thermally inaccessible from the 3MLCT state. A photoproduct of 400 nm excitation of cis-[(bpy)2Ru(NH3)2]2+, cis-[(bpy)2Ru(OH2)2]2+, has been demonstrated to undergo ligand substitution with mononucleotides and was shown to bind readily to double stranded (ds) and single stranded (ss) DNA. The authors predict that the interacting electronic excited states can be tuned by modifying the complex structure to favour photoaquation following visible light excitation. Octahedral polyazine complexes of Rh(III) with a cis-dihalide moiety are also known to undergo ligand substitution upon photolysis with near-UV light.23,29 Morrison and coworkers have studied the DNA photobinding properties of cis[(TL)2RhCl2]+.30,64–66 They demonstrated that [(phen)2RhCl2]+, when irradiated with 308 nm light, photobound to ss- or dsDNA with Fb ≈ 0.006 and 0.001 respectively.30 Dark controls confirm that electronic excitation of the metal complex is required for [(phen)2RhCl2]+ to bind DNA. The authors have also proposed a photooxidation step is necessary to initiate the sequence giving the photosubstitution products.64 Previous assignment of the photolabile state of [(phen)2RhCl2]+ was to the lowest 3 LF state.29 Exchanging phen for methyl substituted 4,7,7,8-tetramethyl-1,10-phenanthroline (Me4phen) or 5,6-dimethyl-1,10-phenanthroline (5,6-Me2phen) had varying impact on the photochemistry.66 The lowest absorption band of [(5,6Me2phen)2RhCl2]+ is red shifted compared to both [(phen)2RhCl2]+ and [(Me4phen)2RhCl2]+. Enhanced photoaquation for [(Me4phen)2RhCl2]+ (Faq = 0.63) compared to [(phen)2RhCl2]+ (Faq = 0.03) when photolysed with 347 nm light was seen. A similar complex, [(phen)(dppz)RhCl2]+ was shown to both photobind and photocleave DNA when irradiated with 311 nm light. The planar dppz ligand is known to facilitate intercalative binding of the metal complex to DNA. The Faq of [(phen)(dppz)RhCl2]+, when irradiated with 355 nm light is reported to be 0.068, reduced compared to [(phen)2RhCl2]+ (Faq = 0.087). Competitive decay to a low lying n → p* state localized on the dppz ligand is possible in the dppz-containing

260

Towards Photodynamic Therapy of Cancer H2N

O HN N N

N

N N

N Ru N

N

HN N N

N

N

N R

O

N

N

N

[(TAP)2Ru(TAP-NH-dGMP)]

N N N

N

N

N Ru N

N N

N N

R

N N

N

N N

N

[(HAT)(phen)Ru(HAT-O-dGMP)]

Figure 8.18 Photoadducts of guanisine monophosphate (dGMP) and different Ru(II) polyazine complexes (TAP = 1,4,5,8-tetraazaphenanthrene, HAT = 1,4,5,8,9,12-hexaazatriphenylene, phen = 1,10-phenanthroline, R = 1′-deoxyribose-5′-phosphate)50,72

complex. The authors noted photocleavage by [(phen)(dppz)RhCl2]+ may occur through a low lying 3IL state. Photobinding DNA through a coordinated ligand. Another mechanism has been implicated in photobinding of the metal complex to DNA through covalent attachment to an ancillary ligand. Moucheron, Kirsch-De Mesmaeker and Kelly have reviewed an extensive series of ruthenium(II) complexes containing polyazaaromatic ligands that are reductively quenched by guanine.50 Excitation of [Ru(TAP)3]2+ , TAP = 1,4,5,8-tetraazaphenanthrene, with visible light gives rise to weak emission from the 3MLCT state. Reductive quenching was observed for dGMP (ket = 2.2 × 109 M−1 s−1), calf thymus (CT) DNA and poly(dG-dC). DNA photoproduct analysis by gel electrophoresis indicated retarded migration of the DNA relative to the nonphotolysed sample. This result is indicative of increased size of the DNA and/or decrease in the overall charge. Change in size and charge would be expected in the event of metal-complex binding to DNA. The metal complex–DNA photoproduct was digested into nucleotides and studied by 1H-NMR spectroscopy and MS. The authors determined the major product was metal complex covalently bound through the 2-carbon of a TAP ligand at the C3-amine of guanine (Figure 8.18). The authors concluded that the complex must bind to the minor groove of DNA to allow for formation of the observed photoadduct. More recently, photoadduct formation during photolysis of DNA with [(phen)Ru(HAT)2]2+ was reported.67 The analogous 2-carbon on the ancillary ligand is again bound to guanine, but at the 5C-oxo site, forming the ether linked photoadduct. The large planar HAT ligands intercalate the DNA base stack in the hydrophobic major groove, lending to preferential formation of the ether linked photoadduct. Design Aspects: Toward Photocleavage of DNA Photoinitiated strand scission of DNA can occur due to excited state electron transfer, hydrogen atom abstraction from a sugar and by reaction of DNA with 1O2.2,52,53,62 The details of these DNA cleavage mechanisms have been outlined above.

Designing Transition Metal Polyazines for Photomodification

261

The ability of an excited state chromophore to oxidize DNA is a reflection of the redox properties of DNA bases and the excited chromophore. Guanine has been identified as the DNA nitrogenous base with the strongest reducing power [E(G/ G+) ≈ 0.92 V versus Ag/AgCl].68–71 When incorporated in dsDNA, oxidation occurs preferentially at the 5′-G of a multi-G stack, suggested to result from a destabilization of the 5′-G donor orbitals, shifting the oxidation potential (E0) of 5′-GGG-3′ to ca. 0.42 V (versus Ag/AgCl).62 Radical cations of the oxidized bases decompose by multiple pathways, including cleavage of the sugar phosphate backbone and base-labile nucleotide oxidation products.52 The chromophore in its ES or an unstable redox state may also be involved in cleavage schemes. Hydrogen atom abstraction from a sugar of DNA by *[(phen)2Rh(phi)]3+ has been proposed to initiate a reaction cascade ending with single-strand scission.58 Early studies have indicated that the 3MLCT states of [Ru(bpy)3]2+ under anoxic and hypotonic conditions are quenched coincidentally with single-strand cleavage of circular plasmid DNA, Frxn = 1.2 × 10−6 when [DNA]/ [Ru(bpy)3]2+ = 9. Recently the photochemistry of the complex [(TAP)2Ru(dppz)]2+ with DNA was reported, showing mixed photochemical pathways.72 The authors noted a mechanism by which the complex photobinds to DNA through a TAP ligand as described above for [Ru(TAP)3]2+. Interestingly, the dppz complex has a strong emission from a long-lived 3MLCT state in room temperature and in aqueous solution (t0 = 1.09 µs). For other Ru(II)-polyazine complexes with the dppz ligand, emission in water is notably weak due to low lying n → p* 3IL state(s) that quenches the emission from the 3MLCT state, possibly through hydrogen bonding of the water to the lone pairs on the nitrogens on the dppz ligand. Intercalation into DNA, however, protects the nitrogens on dppz from water, decreasing quenching of the 3 MLCT state and greatly enhancing emission. Ru(II)-complexes with dppz are often called ‘DNA light switches’.73 The emission of [(TAP)2Ru(dppz)]2+ is efficiently ‘switched off’ upon binding to calf-thymus DNA (t = 60 ns), a very unusual result. Oxygen mediated photocleavage of DNA. Photocleavage of DNA represents a major research effort in the study of DNA photomodifications with application to the development of new PDT agents. Several mechanisms of DNA photocleavage by platinum group metal polyazine complexes have been proposed that require 3O2. DNA photocleavage was reported for [Ru(TL)3]2+, TL = bpy, phen, in aerated solution .74 Mongelli, Brewer and coworkers reported a series of mononuclear Ru(II) complexes containing a potential molecular bridge, [(TL)2Ru(dpp)]2+, where TL = bpy, phen or Ph2phen.75 Enhancement of DNA photocleavage was noted for complexes with more lipophilic terminal ligands, [(Ph2phen)2Ru(dpp)]2+ being the most active, consistent with earlier reports by Barton.56 Hergueta-Bravo, Orellana and coworkers investigated the 3O2 dependence of DNA photocleavage by a series of Ru(II) complexes incorporating the intercalating ligand dibenzo[h,j]dipyrido[3,2a:2′,3′-c]phenazine (ddz).76 The 3MLCT of [(bpy)2Ru(ddz)]2+ state is a less efficient photosensitizer of 1O2 generation relative to [Ru(bpy)3]2+. Enhanced DNA photocleavage by [(bpy)2Ru(ddz)]2+, however, was noted. The increase in DNA photocleavage by [(bpy)2Ru(ddz)]2+ was attributed to strong intercalative binding to the DNA prior to photolysis. A similar observation was made for the complex [(bpy)2Ru(DAP)]+, DAP = 1,12-diazaperylene.77

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The coupling of Ru polyazine chromophores to a cis-PtIICl2 site provides a forum to attach a DNA photocleavage unit to a DNA covalent binding unit. This attachment provides a means to deliver the Ru polyazine chromophore to the DNA target. An example of such a complex, [{(bpy)2Ru(dpp)}2Ru(dpp)PtCl2]6+, binds to DNA through the Pt(II) centre and photocleaves DNA via an oxygen-mediated pathway.78 Oxygen independent photocleavage of DNA. Swavey, Holder and Brewer examined the DNA photocleavage ability of polyazine bridged mixed-metal supramolecules. The supramolecules were constructed with two Ru(II) or two Os(II) light absorber subunits connected to a central cis-RhIIICl2 moiety.42,44 A wide array of supramolecules constructed with a dpp molecular bridge produced nicks in supercoiled circular plasmid DNA with >450 nm light photolysis. The gel showing the photocleavage of DNA by a member of this series, [{(bpy)2Ru(dpp)}2RhCl2]5+, is shown in Figure 8.19. The mechanism of DNA photocleavage is unclear, but this complex functions as a DNA photocleavage agent in the presence or absence of dissolved 3O2. DNA photocleavage is proposed to result from intramolecular ES electron transfer quenching of the 3MLCT state by the cis-RhCl2 center. The resultant metal to metal charge transfer (3MMCT) state has a formally oxidized Ru centre and a formally reduced Rh centre, both of which are capable of mediating DNA cleavage in the absence of 3O2. Importantly, nicks in the DNA backbone caused by most types of photocleavage often are irreparable by the repair enzyme DNA ligase.53

no MC MC MC +hν

N

N II

N

Cl

Ru N N

N

Cl III

N

Rh N N

N

N

N RuII N

N N

N

MC = [{(bpy)2Ru(dpp)}2RhCl2]Cl5

Form II Form I λ

pUC18 DNA

Array of 5W LEDs

Figure 8.19 Photograph of electrophoresis gel showing the efficient photocleavage of pUC18, supercoiled circular plasmid DNA (Form I), with the metal complex [{(bpy)2Ru (dpp)}2RhCl2]Cl5, generating open circular pUC18 DNA (Form II) [bpy = 2,2′-bipyridine, dpp = 2,3-bis(2-pyridyl)pyrazine]. DNA was photolysed with 460 nm centred light from a 5W LED (inset image). From left to right the lanes are: lambda DNA/ HindIII digest molecular weight marker (λ), pUC18 DNA without treatment with metal complex (no MC), pUC18 DNA incubated with metal complex in the dark at 5:1 base pairs to metal complex (MC), and pUC18 DNA photolysed in the presence of metal complex at 5:1 base pairs to metal complex (MC+hν)

Cell Studies with Metal Complexes 263

8.5 Cell Studies with Metal Complexes Cellular uptake and photoinitiated cell death is indicated for several metal polyazinebased light absorbers. Given the short timeframe of this research effort, this is a very promising result, with no strong understanding of means to design systems that will transport into the nucleus. Several groups indicate uptake of the metal complex in the cell to be a function of the ancillary ligands lipophilicity.1,66,79 Metal complexes representing different constructs outlined in preceding sections are promising as PDT agents. Surprisingly few cell studies with Ru, Os or Rh polyazine chromophores have been performed, given the breadth of knowledge of the photochemistry of these complexes with DNA. Cell studies of some Ru, Os and Rh polyazine complexes have revealed their excellent phototoxicity and minimal dark toxicity.33,65,66,79–84 Table 8.2 is a synopsis of the cell studies with transition metal polyazine complexes. Table 8.2 Transition metal polyazine complexes used in cell studies with applications in PDT Complexa [Ru(bpy)3]Cl2 and [Ru(phen)3]Cl2 [(bpy)2Ru(ddz)]Cl2 [(TL)2Ru(dppz)]Cl2TL = bpy, phen, mcmb, Ph2phen

cis-[(TL)2RhCl2]Cl TL = phen, 5,6-Me2phen, Me4phen, Ph2phen cis-[(phen)(dppz)RhCl2]Cl

[(phen)(chrysi)Rh(dmb(arg)8-DYE)](O2CCF3)11 cis-[Rh2(µ-O2CMe)2(dppz) (bpy)](O2CMe)2 [{(bpy)2M(dpp)}2RhCl2]Cl5, M = Ru or Os

Description of Interaction

Ref.

• Cellular uptake following photolysis • Phototoxic at concentrations >10−4 M • Selective uptake and luminescence enhancement by nonviable cells • Cell photokilling not investigated • Cellular uptake as a function of lipophilicity, size and overall charge • Faster uptake for complexes with lipophilic ligands (e.g. TL = Ph2phen) • Cell photokilling not investigated • Uptake investigated as function of lipophilicity • Uptake observed only for TL = Me4phen • Phototoxic to tumour cell lines at 55 µM and irradiation with >400 nm light • Penetrates cellular membranes and viral coats • Phototoxic to tumour cells at 40 µM and irradiation with 311 nm light • Deactivates viral DNA within intact virus • Metal complex with octa(arginine) tail and fluorescent dye • Rapid uptake observed at 5 µM • Cell photokilling not investigated • Dark toxicity reported LC50 = 208 µM • Phototoxic LC50 = 44 µM with 30 min, 400– 700 nm irradiation • No dark toxicity observed (tested up to 120 µM) • Phototoxic at 5 µM, with 4 min , 400–1000 nm irradiation

80 81

79

66

65

82

33,83

84

a bpy = 2,2′-bipyridine; phen = 1,10-phenanthroline; mcmb = 4-carboxy-4′-methyl-2,2′-bipyridine, Ph2phen = 4,7diphenyl-1,10-phenanthroline; dppz = dipyrido[3,2-a:2′,3′-c]phenazine; 5,6-Me2phen = 5,6-dimethyl-1,10-phenanthroline; Me4phen = 3,4,7,8-tertramethyl-1,10-phenanthroline; chrysi = a-chrysenequinone diimine; ddz = dibenzo [h,j]dipyrido[3,2-a:2′,3′-c]phenazine; dpp = 2,3-bis(2-pyridyl)pyrazine.

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8.5.1 Cellular Uptake and Phototoxicity of Transition Metal Polyazine Complexes Recent studies of the interaction of platinum group polyazine complexes with cells indicate that uptake of metal complexes by cells is greater with a compromised cellular envelope. Dobrucki investigated [Ru(bpy)3]Cl2 and [Ru(phen)]Cl2 as confocal microscopy imaging dyes of J144 mouse macrophages and normal human fibroblasts (NFB).80 The metal complexes were internalized by the cells with endocytosis. Neither metal complex penetrated intact plasma membranes under dark conditions. Visible light photolysis of cells with internalized metal complex resulted in redistribution of the observed luminescence within the cell. Dobrucki suggests that Type II photooxidation of the cell membrane compromises its integrity, allowing passage of the metal complexes. Photoinduced cell death was noted for concentrations of [Ru(phen)3]2+ greater than 0.2 mM and irradiation with 490 nm light. JimenezHernandez, Orellana, et al. studied [(bpy)2Ru(ddz)]2+ as a selective stain of nonviable rat hepatocytes.81 A commonly used technique to image nonviable cells with fluorescence microscopy uses intercalation of the ethidium homodimer ion into DNA, which greatly enhances ethidium emission. This provides red fluorescent imaging of the nonviable cells. The planar aromatic ddz ligand of [(bpy)2Ru(ddz)]2+ readily intercalated DNA, also exhibiting a potent DNA light switch effect. Upon passing through the leaky membrane of dead cells, [(bpy)2Ru(ddz)]2+ is rapidly bound by nuclear DNA to give an emissive label of nonviable cells. A metal complex bound to an octa(arginine) tether [(arg)8] exhibited enhanced cellular uptake and association with nuclear DNA.82 Brunner and Barton studied the cellular uptake of [(phen)(chrysi)Rh{dmb-(arg)8-DYE}](O2CCF3)11, where DYE is a fluorescein or thiazole orange fluorescent tag. Uptake of the fluorescein dyelabelled metal complex was studied by confocal microscopy of metal complex bound HeLa cells. Confocal microscopy images of cells incubated with 5 µM showed that the dye was rapidly incorporated into the nucleus. The authors suggest that the design of the studied complexes could be adapted to build and understand the cellular uptake mechanisms of other metal complexes. Mononuclear ruthenium(II) and rhodium(III) polyazine complexes exhibit cellular uptake mechanisms related to the lipophilicity of the metal complex. Puckett and Barton studied the cellular uptake of a series of [(TL)2Ru(dppz)]Cl2 complexes, with TL = bpy, phen, Ph2phen, and 4-carboxy-4′-methyl-2,2′-bipyrindine (mcmb).79 Uptake was observed using confocal microscopy with direct excitation of the Ruex centred chromophore λmax = 488 nm . The complex [(Ph2phen)Ru(dppz)]Cl2 displayed the largest mean luminescence intensity inside the cell. The authors related uptake to the octanol/water partition coefficient of the metal complexes, suggesting a passive transport mechanism. Minimal emission was observed within the nucleus, suggesting that transport across the nuclear envelop is retarded. Loganathan and Morrison reported a similar enhanced cellular uptake with increasing lipophilicity for cis-[(TL)2RhCl2]Cl as a function of TL methylation.66 The authors report that [(Me4phen)2RhCl2]Cl, at 55 µM and with >400 nm light irradiation, caused a 20% reduction in cell growth relative to dark controls. The authors were unable to assess the mechanism of cell killing, but noted [(Me4phen)2RhCl2]Cl to be an efficient photobinder of DNA. Another DNA photobinding complex, [(phen)(dppz)RhCl2]Cl, was reported to be phototoxic at 40 µM with 311 nm irradiation.65 This complex was

Cell Studies with Metal Complexes 265

also shown to deactivate the DNA of a virus, indicating transport through intact viral envelope. 8.5.2 Polymetallic Complex Phototoxicity Studies Four recent examples of multimetallic polyazine complexes, in two separate studies, were reported to have a dramatic increase in toxicity upon photolysis. Angeles-Boza, Dunbar, Turro and coworkers studied the dirhodium(II) complexes, cis-[Rh2(m-O2 CMe)2(dppz)(TL)](O2CMe)2, where TL = MeOH and MeCOOH; dppz; or bpy.33,83 There was little enhancement of toxicity toward human skin fibroblasts upon photolysis in the presence of cis-[Rh2(m-O2CMe)2(dppz)(MeOH)(h1- O2CMe)](O2CMe). The dark LC50 values for cis-[Rh2(m-O2CMe)2(dppz)2](O2CMe)2 and cis-[Rh2(m-O2 CMe)2(dppz)(bpy)](O2CMe)2 were 140 µM and 210 µM, respectively. This is slightly less toxic than cisplatin (LC50 = 131 µM) and much less toxic than hematoporphyrin under the same conditions (LC50 = 20 µM). Each complex displayed comparable phototoxicity (LC50 ≈ 40 µM) with >395 nm light irradiation for 15 min. The complex cis-[Rh2(m-O2CMe)2(dppz)(bpy)](O2CMe)2 had the greatest difference between light and dark toxicity. Holder, Storrie, Brewer et al. investigated the phototoxicity of mixed-metal supramolecules of the form [{(bpy)2M(dpp)}2RhCl2]Cl5, where M = Ru or Os.84 No appreciable dark toxicity was observed in Vero cells for any concentration of either metal complex, up to 120 µM. Excess metal complex was washed from the cells prior to photolysis. Irradiation with > 460 nm for 5 min resulted in dramatic decrease in cell viability (Figure 8.20). As the concentration of [{(bpy)2Ru(dpp)}2RhCl2]Cl5 in N

N RuII N

N N

N

Cl

Cl

N

RhIII N

N N

N

N RuII N

N N

N

MC = [{(bpy)2Ru(dpp)}2RhCl2]Cl5 Cells treated with MC

illumination area

growth

after 48 hr growth period

calcein AM imaging of live cells

ethidium imaging of dead cells

Figure 8.20 (Plate 11) Micrographs of Vero cells pretreated with [{(bpy)2Ru(dpp)}2RhCl2]C l5, rinsed, and illuminated with 400–1000 nm light showing the high level of light-activated cell killing [bpy = 2,2′-bipyridine, dpp = 2,3-bis(2-pyridyl)pyrazine]. (From left to right) immediately after photolysis (light exposure within circle); after 48 hour growth period; live cell (green) visualized with calcein AM fluorescent dye; dead cell (red) visualized with ethidium homodimer-1 fluorescent dye84 (See colour plate section)

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media used to pre-treat the cells was increased from 5 to 120 µM, the number of viable cells following photolysis approached zero. The Os complex, [{(bpy)2Os(dpp)}2 RhCl2]Cl5, exhibited decreased phototoxicity relative to the Ru complex, with the ratio of cells present before and after photolysis decreasing toward unity as a function metal complex concentration. The difference in phototoxicity is quite surprising given the similar DNA photoreactivity. This may be due to the lifetime of the reactive state of each complex, with the Os systems providing shorter ES lifetimes. The cells studies of these metal complexes show great promise. Neither study of multimetallic complexes could comment on the mechanism of cell killing. Both report the photosensitizers of interest to be efficient DNA photocleavage agents.33,43,44,83

8.6 Conclusions and Future Directions Platinum group metal polyazine complexes have pronounced DNA photochemistry and show unusual promise as photodynamic therapy (PDT) agents. Recent discoveries contribute to the understanding of the electronic excited state interactions of these systems with DNA. Studies of photoactivated interactions of platinum group complexes with biomolecules also provide insight into the basic photophysics of these metal complexes. DNA can be a good probe of the excited state dynamics of metal complexes. Polyazine complexes of transition metal centres, especially ruthenium and osmium, exhibit strong light absorbing properties in the UV and in the visible region, with interesting photoreactivity. Excitation populates reactive electronic excited states with properties very different than the ground state molecules. Ruthenium(II) polyazine complexes are noted visible light absorbers with typically long-lived Ru(dp) → dpp(p*) charge transfer (3MLCT) states known to photosensitize 1O2. Osmium(II) polyazine complexes have similar light absorbing properties to ruthenium(II), but with electronic transitions shifted to lower energy. The phosphorescence of 3MLCT states provides a convenient handle to reactivity of these 3MLCT states. Rhodium(III) polyazine complexes, though poor visible light absorbers, have interesting reactive internal ligand (3IL) and ligand field (3LF) electronic excited states that photocleave and photobind DNA. The studies to date provide significant motivation for continued study of Rh(III) complexes. Polyazine complexes incorporating dirhodium(II) centres possess electronic excited states arising from the delocalized metal–metal(dp*) highest occupied molecular orbital, giving them potent photochemical reactivity. Polyazine bridged complexes of ruthenium(II) and rhodium(III) are both visible light absorbers and possess the reactive Rh(III) centre. Systems can be designed to have a lowest lying Ru(dp) → Rh(dσ*) charge transfer (3MMCT) state that is reactive with, among other substrates, DNA. Structural modifications of these assemblies provide a unique forum to modulate properties. The forum of photomodification of DNA by metal polyazine complexes has been developing over the last three decades. Researchers in the field have discovered multiple modes of photochemical action, including metal complex mediated

Acknowledgements

267

photooxidation of DNA bases, photobinding metal polyazine chromophores to DNA and photocleavage of the DNA backbone by the excited inorganic chromophore. These studies show both the complexity of the interactions of metal complexes with DNA in the excited state and the potential promise of these complexes in PDT. The excited states of metal polyazine complexes are easily tuned by modification of the ligand environment, as is tuning of ground state interactions of these systems with DNA. Metal complexes that associate with DNA to a greater extent prior to photolysis tend to have enhanced efficiency of DNA photomodification. Given the typically low concentration of DNA, metal complex–DNA interactions are key to photoreaction efficiency. Systems incorporating extended planar, aromatic, polyazine ligands or systems containing lipophilic polyazine ligands generally interact strongly with DNA. The studies of metal complexes and their light activated modification of DNA show the promise of metal complexes in this forum. The studies summarized herein illustrate examples of a variety of photoreactions with DNA. Many systems react with DNA by yet to be understood mechanisms. Surprisingly few studies of platinum group metal polyazine complexes with cells have been undertaken and it is clear that much is to be learned in this forum. The utility of metal complexes as chromophores in photoinitiated cell killing schemes remains clear. The photochemical mechanisms of platinum group metal polyazine complexes leading to cell death have yet to be explored. The more detailed studies of DNA photomodification noted for these systems suggest this as a possible mechanism of cell photokilling. Cellular uptake mechanisms of metal polyazine complexes are currently being investigated to understand the relationship between ligand lipophilicity and uptake. While Barton and coworkers show that rational design strategies can be used to develop platinum group metal polyazine complexes which readily enter cells, many surprises are being uncovered. Studies of multinuclear polyazine complexes illustrate the utility of this class of photosensitizer in future photodynamic therapy schemes. The low dark toxicity and high light activated cell killing makes these supramolecular systems particularly promising to develop new, potent PDT agents. Continuing studies within the field of platinum group metal polyazine complexes as PDT agents, will ultimately lead to their clinical use in the treatment of cancer.

Acknowledgements Special thanks to Prof. Brenda S.J. Winkel for her frequent questions, discussions and collaborations on studies of mixed-metal supramolecules and their photochemistry with DNA. The authors would especially like to thank the National Science Foundation for their generous support of this work (CHE-0408445). The authors thank Theralase, Inc. for continued collaboration to investigate the PDT action of supramolecular complexes. The authors would also like to thank Dr Shamindri M. Arachchige, Dr Krishnan Rangan, Avijita Jain, Samantha Hopkins and Anne-Marie C. Overstreet for help with this chapter.

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Abbreviations 5,6-Me2phen bpm bpy DAP ddz dpp dppz dsdp EA ED en ES et f GS HAT HOMO ic IL isc kx LA *LA LF LUMO Me4phen MLCT MMCT MMLCT nr p PDT Ph2phen phen phi q Q rxn TAP tpy Fx

5,6-dimethyl-1,10-phenanthroline 2,2′-bipyrimidine 2,2′-bipyridine 1,12-diazapyrylene dibenzo[h,j]dipyrido[3,2-a:2′,3′-c]phenazine 2,3-bis(2-pyridyl)pyrazine dipyrido[3,2-a:2′,3′-c]phenazine 4,7-di(3-sulfurylphenyl)-1,10-phenanthroline electron acceptor electron donor excited state energy transfer excited state light absorber excited state electron transfer fluorescence ground state light absorber 1,4,5,8,9,12-hexaazatriphenylene highest occupied molecular orbital internal conversion internal ligand intersystem crossing rate constant of process ‘x’ ground state light absorber excited state light absorber ligand field lowest unoccupied molecular orbital 3,4,7,8-tetramethyl-1,10-phenanthroline metal to ligand charge transfer metal to metal charge transfer metal–metal to ligand charge transfer nonradiative decay phosphorescence photodynamic therapy 4,7-diphenyl-1,10-phenanthroline 1,10-phenanthroline 9,10-phenanthrenequinone diimine bimolecular deactivation quencher photochemical reaction 1,4,5,8-tetraazaphenanthrene 2,2′:6′,2″-terpyridine quantum yield of process ‘x’

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References 1. Ali, H.; van Lier, J.E.; Metal complexes as photo- and radiosensitizers; Chem. Rev., 1999, 99, 2379–2450. 2. Kirsch-De Mesmaeker, A.; Lecomte, J.-P.; Kelly, J.M.; Photoreactions of metal complexes with DNA, especially those involving a primary photo-electron transfer; Top. Curr. Chem.; 1996, 177, 25–76. 3. Clarke, M.J.; Non-platinum chemotherapeutic metallopharmaceuticals; Chem. Rev., 1999, 99, 2511–2533. 4. Clarke, M.J.; Ruthenium metallopharmaceuticals; Coord. Chem. Rev., 2003, 236, 209–233. 5. Erkkila, K.E.; Odom, D.T.; Barton, J.K.; Recognition and reaction of metallointercalators with DNA; Chem. Rev., 1999, 99, 2777–2795. 6. McMillin, D.R.; McNett, K.M.; Photoprocesses of copper complexes that bind to DNA; Chem. Rev., 1998, 98, 1201–1219. 7. Stochel, G.; Wanat, A.; Kulis, E.; Stasicka, Z.; Light and metal complexes in medicine; Coord. Chem. Rev., 1998, 171, 203–220. 8. Szacilowski, K.; Macyk, W.; Drzewiecka-Matuszek, A.; Brindell, M.; Stochel, G.; Bioinorganic photochemistry: frontiers and mechanisms; Chem. Rev., 2005, 105, 2647– 2694. 9. Castano, A.P.; Demidova, T.N.; Hamblin, M.R.; Mechanisms in photodynamic therapy: part one – photosensitizers, photochemistry and cellular localization; Photodiagn. Photodyn. Ther., 2004, 1, 279–293. 10. Castano, A.P.; Demidova, T.N.; Hamblin, M.R.; Mechanisms in photodynamic therapy: part two – cellular signaling, cell metabolism, and modes of cell death; Photodiagn. Photodyn. Ther., 2005, 2, 1–23. 11. Castano, A.P.; Demidova, T.N.; Hamblin, M.R.; Mechanisms in photodynamic therapy: part three – photosensitizer pharmicokinetics, biodistribution, tumor localization and modes of tumor destruction; Photodiagn. Photodyn. Ther., 2005, 2, 91–106. 12. Allison, R.R.; Mota, H.C.; Sibata, C.H.; Clinical Pd/Pdt in North America: An historical review; Photodiagn. Photodyn. Ther., 2004, 1, 263–277. 13. Kessel, D.; Photodynamic therapy: from the beginning; Photodiagn. Photodyn. Ther., 2004, 1, 3–7. 14. Allison, R.R.; Downie, G.H.; Cuenca, R.; Hu, X.-H.; Childs, C.J.H.; Sibata, C.H.; Photosensitizers in clinical PDT; Photodiagn. Photodyn. Ther., 2004, 1, 27–42. 15. Triesscheijn, M.; Baas, P.; Schellens, J.H.M.; Stewart, F.A.; Photodynamic therapy in oncology; Oncologist, 2006, 11, 1034–1044. 16. Englman, R.; Jortner, J.; The energy gap law for radiationless transitions in large molecules; Mol. Phys., 1970, 18, 145–164. 17. Calladine, C.R.; Understanding DNA: The Molecule and How It Works, 3rd Edn, Elsevier Academic Press, San Diego, CA, 2004. 18. Kalyanasundaram, K.; Photochemistry of Polypyridine and Porphyrin Complexes, Academic Press, San Diego, CA, 1992. 19. Juris, A.; Balzani, V.; Barigletti, F.; Campagna, S.; Belser, P.; Von Zelewsky, A.; Ru(II) polypyridine complexes photophysics, photochemistry, electrochemistry, and chemiluminescence; Coord. Chem. Rev., 1988, 84, 85–277. 20. Balzani, V.; Moggi, L.; Scandola, F.; In Supramolecular Photochemistry; Balzani, V., Ed., D. Reidel Publishing Company, 1987, 1–28. 21. Demas, J.N.; Taylor, D.G.; Intersystem crossing yields in ruthenium(II) and osmium(II) photosensitizers; Inorg. Chem., 1979, 18, 3177–3179. 22. Singh, T.N.; Turro, C.; Photointiated DNA binding by Cis-[Ru(Bpy)2(Nh3)2]2+; Inorg. Chem., 2004, 43, 7260–7262. 23. Ford, P.C.; Wink, D.; Dibenedetto, J.; In Progress in Inorganic Chemistry, Vol. 30, Lippard, S.J., Ed., John Wiley & Sons, Inc., New York, 1983,213–271.

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24. Kober, E.M.; Caspar, J.V.; Lumpkin, R.S.; Meyer, T.J.; Application of the energy gap law to excited-state decay of osmium(II)-polypyridine complexes: calculation of relative nonradiative decay rates from emission spectral profiles; J. Phys. Chem., 1986, 90, 3722–3734. 25. Caspar, J.V.; Kober, E.M.; Sullivan, B.P.; Meyer, T.J.; Application of the energy gap law to the decay of charge-transfer excited states; J. Am. Chem. Soc., 1982, 104, 630–632. 26. Crosby, G.A.; Elfring, W.H., Jr.; Excited states of mixed ligand chelates of ruthenium(II) and rhodium(III); J. Phys. Chem., 1976, 80, 2206–2211. 27. Demas, J.N.; Crosby, G.A.; Quantum efficiencies of transition-metal complexes.I. d-d Luminescence; J. Am. Chem. Soc., 1970, 92, 7262–7270. 28. Crosby, G.A.; Spectroscopic investigations of excited states of transition-metal complexes; Acc. Chem. Res., 1975, 8, 231–238. 29. Muir, M.M.; Huang, W.-L.; Photoaquation of some complexes of rhodium(III); Inorg. Chem., 1973, 12, 1831–1835. 30. Mahnken, R.E.; Billadeau, M.A.; Nikonowicz, E.P.; Morrison, H.; Toward the development of photo cis-platinum reagents. Reaction of cis-dichlorobis(1,10-phenanthroline)rh odium(III) with calf thymus DNA, nucleotides, and nucleosides; J. Am. Chem. Soc., 1992, 114, 9253–9265. 31. Sherman, S.E.; Lippard, S.J.; Structural aspects of platinum anticancer drug interactions with DNA; Chem. Rev., 1987, 87, 1153–1181. 32. Bradley, P.M.; Fu, P.K.-L.; Turro, C.; Excited state properties of Rh2(O2CCH3)4: solution photochemistry and photoinitiated DNA cleavage; Comments Inorg. Chem., 2001, 22, 393–426. 33. Angeles-Boza, A.M.; Bradley, P.M.; Fu, P.K.-L.; Wicke, S.E.; Bacsa, J.; Dunbar, K.R.; Turro, C.; DNA binding and photocleavage in vitro by new dirhodium(II) dppz complexes: correlation to cytotoxicity and photocyotoxicity; Inorg. Chem., 2004, 43, 8510–8519. 34. De Cola, L.; Belser, P.; Photoinduced energy and electron transfer processes in rigidly bridged dinuclear Ru/Os complexes; Coord. Chem. Rev., 1998, 177, 301–346. 35. Furue, M.; Hirata, M.; Kinoshita, S.; Kushida, T.; Kamachi, M.; Intramolecular electrontransfer of covalently-linked polypyridine ruthenium(II)/rhodium(III) binuclear complexes in the excited state. observation of the Marcus inverted region; Chem. Lett., 1990, 2065–2069. 36. Indelli, M.T.; Bignozzi, C.A.; Harriman, A.; Schoonover, J.R.; Scandola, F.; Four intercomponent processes in a Ru(II)-Rh(III) polypyridine dyad: electron transfer from excited donor, electron transfer to excited acceptor, charge recombination, and electronic energy transfer; J. Am. Chem. Soc., 1994, 116, 3768–3779. 37. Kalyanasundaram, K.; Gratzel, M.; Nazeeruddin, M.K.; Excited-state interactions in ligand-based chromophore-quencher complexes containing rhodium(III) and ruthenium(II) polypyridyl units; J. Phys. Chem., 1992, 96, 5865–5872. 38. Kalyanasundaram, K.; Nazeeruddin, M.K.; Photophysics and photoredox reactions of ligand-bridged binuclear polypyridyl complexes of ruthenium(II) and their monomeric analogues; Inorg. Chem., 1990, 29, 1888–1897. 39. Kleverlaan, C.J.; Indelli, M.T.; Bignozzi, C.A.; Pavanin, L.; Scandola, F.; Hasselman, G.M.; Meyer, G.J.; Stepwise charge separation in heterotriads. binuclear Ru(II)-Rh(III) complexes on nanocrystalline titanium dioxide; J. Am. Chem. Soc., 2000, 122, 2840–2849. 40. Fuchs, Y.; Lofters, S.; Dieter, T.; Shi, W.; Morgan, R.; Strekas, T.C.; Gafney, H.D.; Baker, A.D.; Spectroscopic and electrochemical properties of dimeric ruthenium(II) diimine complexes and determination of their excited state redox properties; J. Am. Chem. Soc.; 1987, 109, 2691–2697. 41. Molnar, S.M.; Jensen, G.E.; Volger, L.M.; Jones, S.W.; Laverman, L.; Bridgewater, J.S.; Richter, M.M.; Brewer, K.J.; Photochemical properties of mixed-metal supramolecular complexes; J. Photochem. Photobiol. A, 1994, 80, 315–322. 42. Swavey, S.; Brewer, K.J.; Synthesis and study of Ru,Rh,Ru triads: modulation of orbital energies in a supramolecular architecture; Inorg. Chem.; 2002, 41, 4044–4050. 43. Swavey, S.; Brewer, K.J.; Visible light induced photocleavage of DNA by a mixed-metal supramolecular complex: [{(bpy)2Ru(dpp)}2RhCl2]5+; Inorg. Chem., 2002, 41, 6196–6198.

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44. Holder, A.A.; Swavey, S.; Brewer, K.J.; Design aspects for the development of mixedmetal supramolecular complexes capable of visible light induced photocleavage of DNA; Inorg. Chem., 2004, 43, 303–308. 45. Marcus, R.A.; Chemical and electrochemical electron transfer theory; Annu. Rev. Phys. Chem.; 1964, 15, 155–196. 46. Foote, C.S. Definition of Type I and Type II Photosensitized Oxidation; Photochem. Photobiol. 1991, 54, 659. 47. Forster, T.; Transfer mechanisms of electronic excitation; Discuss. Faraday Soc., 1959, 27, 7–17. 48. Dexter, D.L.; A theory of sensitized luminescence in solids; J. Chem. Phys., 1953, 21, 836–850. 49. Demas, J.N.; Diemente, D.; Harris, E.W.; Oxygen quenching and charge-transfer excited states of ruthenium(II) complexes. Evidence for singlet oxygen production; J. Am. Chem. Soc., 1973, 95, 6864–6865. 50. Moucheron, C.; Kirsch-De Mesmaeker, A.; Kelly, J.M.; Photoreactions of ruthenium(II) and osmium(II) complexes with deoxyribonucleic acid (DNA); J. Photochem. Photobiol. B, 1997, 40, 91–106. 51. Zeglis, B.M.; Pierre, V.C.; Barton, J.K.; Metallo-intercalators and metallo-insertors; J. Chem. Soc. Chem. Commun., 2007, 4565–4579. 52. Armitage, B.; Photocleavage of nucleic acids; Chem. Rev., 1998, 98, 1171–1200. 53. Pogozeleski, W.K.; Tullius, T.D.; Oxidative strand scission of nucleic acids: Routes initated by hydrogen abstraction from the sugar moiety; Chem. Rev., 1998, 98, 1098–1107. 54. Vinograd, J.; Lebowitz, J.; Physical and topological properties of circular DNA; J. Gen. Physiol., 1966, 49, 103–125. 55. van der Maarel, J.R.C.; Zakharova, S.S.; Jesse, W.; Backendorf, C.; Egelhaaf, S.U.; Lapp, A.; Supercoiled DNA; Plectonemic structure and liquid crystal formation; J. Phys. Condens. Matter, 2003, 15, S183–S189. 56. Kumar, C.V.; Barton, J.K.; Turro, N.J.; Photophysics of ruthenium complexes bound to double helical DNA; J. Am. Chem. Soc. 1985, 107, 5518–5523. 57. Mei, H.Y.; Barton, J.K.; Chiral probe for A-form helices of DNA and RNA: tris(tetrame thylphenanthroline)ruthenium(II); J. Am. Chem. Soc., 1986, 108, 7414–7416. 58. Sitlani, A.; Long, E.C.; Pyle, A.M.; Barton, J.K.; DNA photocleavage by phenanthrene diimine complexes of rhodium(III): Shape selective recognition and reaction; J. Am. Chem. Soc., 1992, 114, 2303–2312. 59. Barton, J.K.; Metals and DNA: Molecular left-handed compliments; Science, 1986, 233, 727–734. 60. Manning, G.S.; Ray, J.; Counterion condensation revisted; J. Biomol. Struct. Dyn., 1998, 16, 461–476. 61. Junicke, H.; Hart, J.; Kisko, J.; Glebov, O.; Kirsch, I.R.; Barton, J.K.; Bioinorganic chemistry special feature: A rhodium complex for high-affinity DNA base-pair mismatch recognition; Proc. Natl. Acad. Sci. USA, 2003, 100, 3737–3742. 62. Burrows, C.J.; Muller, J.G.; Oxidative nucleobase modifications leading to strand scission; Chem. Rev., 1998, 98, 1109–1151. 63. Poklar, N.; Pilch, D.S.; Lippard, S.J.; Redding, E.A.; Dunham, S.U.; Breslauer, K.; Influence of cisplatin intrastrand crosslinking on the conformation, thermal stability, and energetics of a 20-mer DNA duplex; Proc. Natl. Acad. Sci. USA, 1996, 93, 7606–7611. 64. Morrison, H.; Harmon, H.; ‘Hot spots’ associated with the photoinduced binding of cisdichloro bis(1,10-phenanthroline)rhodium(III) chloride to HIV-1 and C-Raf DNA; Photochem. Photobiol., 2000, 72, 731–738. 65. Menon, E.L.; Perera, R.; Navarro, M.; Kuhn, R.J.; Morrison, H.; Phototoxicity against tumor cells and sindbis virus by an octahedral rhodium bisbipyridyl complex and evidence for the genome as a target in viral photoinactivation; Inorg. Chem., 2004, 43, 5373–5381. 66. Loganathan, D.; Morrison, H.; Effect of ring methylation on the photophysical, photochemical and photobiological properties of cis-dichlorobis(1,10-phenanthrolin)rhodium (III)chloride; Photochem. Photobiol., 2006, 82, 237–247.

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67. Blasiu, R.; Nierengarten, H.; Luhmer, M.; Constant, J.-F.; Defrancq, E.; Dumy, P.; van Dorsselaer, A.; Moucheron, C.; Kirsch-De Mesmaeker, A.; Photoreaction of [Ru(HAT)2phen]2+ with guanosine-5′-monophosphate and DNA: Formation of new types of photoadducts; Chem. Euro.J., 2005, 11, 1507–1517. 68. Kawanishi, S.; Oikawa, S.; Murata, M.; Tsukitome, H.; Saito, I.; Site-specific oxidation at GG and GGG sequences in double-stranded DNA by benzoyl peroxide as a tumor promoter; Biochemistry, 1999, 38, 16733–16739. 69. Prat, F.; Houk, K.N.; Foote, C.S.; Effect of guanine stacking on the oxidation of 8-oxoguanine in B-DNA; J. Am. Chem. Soc., 1998, 120, 845–846. 70. Saito, I.; Takayama, M.; Sugiyama, H.; Nakatani, K.; Photoinduced DNA cleavage via electron transfer: Demonstration that guanine residues located 5′ to guanine are the most electron-donating sites; J. Am. Chem. Soc., 1995, 117, 6406–6407. 71. Sugiyama, H.; Saito, I.; Theoretical studies of GG-specific photocleavage of DNA via electron transfer: Significant lowering of ionization potential and 5′-localization of homo of stacked GG bases in B-form DNA; J. Am. Chem. Soc., 1996, 118, 7063–7068. 72. Ortmans, I.; Elias, B.; Kelly, J.M.; Moucheron, C.; Kirsch-De Mesmaeker, A.; [Ru(TAP)2(dppz)]2+: A DNA intercalating complex, which luminesces strongly in water and undergoes photo-induced proton-coupled electron transfer with guanosine-5′monophosphate; Dalton Trans., 2004, 668–676. 73. Friedman, A.E.; Chambron, J.-C.; Sauvage, J.-P.; Turro, N.J.; Barton, J.K.; Molecular ‘light switch’ for DNA; Ru(bpy)2(dppz)2+; J. Am. Chem. Soc., 1990, 112, 4960–4692. 74. Fleisher, M.B.; Waterman, K.C.; Turro, N.J.; Barton, J.K.; Light-induced cleavage of DNA by metal complexes; Inorg. Chem., 1986, 25, 3549–3551. 75. Mongelli, M.T.; Heinecke, J.; Mayfield, S.; Okyere, B.; Winkel, B.S.J.; Brewer, K.J.; Variation of DNA photocleavage efficiency for {(TL)2Ru(dpp)]Cl2 complexes where TL = 2,2′bipyridine, 1,10-phenanthroline, or 4,7-diphenyl-1,10-phenanthroline; J. Inorg. Biochem., 2006, 100, 1983–1987. 76. Hergueta-Bravo, A.; Jimenez-Hernandez, M.E.; Montero, F.; Oliveros, E.; Orellana, G.; Singlet-oxygen mediated DNA photocleavage with Ru(II) polypyridyl complexes; J. Phys. Chem. B, 2002, 106, 4010–4017. 77. Chouai, A.; Wicke, S.E.; Turro, C.; Bacsa, J.; Dunbar, K.R.; Wang, D.; Thummel, R.P.; Ruthenium(II) complexes of 1,12-diazaperylene and their interactions with DNA; Inorg. Chem., 2005, 44, 5996–6003. 78. Miao, R.; Mongelli, M.T.; Zigler, D.F.; Winkel, B.S.J.; Brewer, K.J.; A multifunctional tetrametallic Ru-Pt supramolecular complex exhibiting both DNA binding and photocleavage; Inorg. Chem., 2006, 45, 10413–10415. 79. Puckett, C.A.; Barton, J.K.; Methods to explore cellular uptake of ruthenium complexes; J. Am. Chem. Soc., 2007, 129, 46–47. 80. Dobrucki, J.W.; Interaction of oxygen-sensitive luminescent probes Ru(phen)32+ and Ru(bipy)32+ with animal and plant cells in vitro. Mechanism of phototoxicity and conditions for non-invasive oxygen measurements; J. Photochem. Photobiol. B, 2001, 65, 136–144. 81. Jimenez-Hernandez, M.E.; Orellana, G.; Montero, F.; Portoles, M.T.; A ruthenium probe for cell viability measurement using flow cytometry, confocal microscopy and timeresolved luminescence; Photochem. Photobiol., 2000, 72, 28–34. 82. Brunner, J.; Barton, J.K.; Targeting DNA mismatches with rhodium intercalators functionalized with a cell-penetrating peptide; Biochemistry, 2006, 45, 12295–12302. 83. Angeles-Boza, A.M.; Bradley, P.M.; Fu, P.K.-L.; Shatruk, M.; Hilfiger, M.G.; Dunbar, K. R.; Turro, C.; Phototoxicity of a new Rh2(II,II) complex: Increase in cytotoxicity upon irradiation similar to that of Pdt agent hematoporphyrin; Inorg. Chem., 2005, 44, 7262–7264. 84. Holder, A.A.; Zigler, D.F.; Tarrago-Trani, M.T.; Storrie, B.; Brewer, K.J.; Photobiological impact of [{(bpy)2Ru(dpp)}2RhCl2]Cl5 and [{(bpy)2Os(dpp)}2rhcl2]Cl5 [bpy = 2,2′-bipyridine; dpp = 2,3-bis(2-pyridyl)pyrazine] on Vero cells; Inorg. Chem., 2007, 46, 4760–4762.

9 Platinated Oligonucleotides: Synthesis and Applications for the Control of Gene Expression Vicente Marchán and Anna Grandas

9.1 Therapeutic Applications of Synthetic Oligonucleotides The genetic information necessary for making a living organism is encoded in DNA (deoxyribonucleic acid), a very large biopolymer associated with many copies of several proteins in chromosomes. The individual units of DNA that contain the instructions for producing proteins are called genes. However, the flow of information from DNA to proteins is mediated by a different nucleic acid, mRNA (messenger ribonucleic acid), which differs from DNA in that deoxyribose is replaced by ribose and thymine by uracil. According to the central tenet of molecular biology, the information encoded by DNA in genes is transcribed into information in RNA (although in some cases, such as in retroviruses, information flows from RNA to DNA), which in turn is translated into the functional protein. The availability of molecules that specifically target either protein or DNA or RNA sequences is essential for the treatment of diseases of genetic, viral or bacterial origin and malignant tumours. In this respect, the extraordinary capacity for recognition of nucleic acids through hydrogen bonding between complementary bases may allow the use of nucleotide-based molecules to control the expression of a particular gene. In 1978 Zamecnik and Stephenson reported for the first time the use of small nucleic acid fragments, called antisense oligonucleotides (AS-O), as therapeutic Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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transcription

action may be blocked by complex formation with synthetic oligonucleotides (aptamers)

translation

dsDNA

mRNA

protein

blocked by antigene oligonucleotides.

blocked by antisense oligonucleotides, ribozymes, siRNAs.

Triplex formation:

Duplex formation:

antisense oligonucleotides

ribozymes

Figure 9.1 Use of synthetic oligonucleotides for the control of gene expression

agents that could inhibit gene expression.1 Antisense technology is based on the specific hybridization of synthetic single-stranded oligonucleotides (some 15–20 bases in length) to a complementary mRNA sequence through Watson–Crick pairing, which prevents the expression of the target gene2,3 (Figure 9.1). Unlike conventional drugs, which bind proteins and modulate their functions, antisense oligonucleotides act at the mRNA level, inhibiting protein biosynthesis through two major mechanisms. When a highly stable heteroduplex is formed between AS-O and mRNA, the ribosomal machinery cannot process the information carried by that mRNA, resulting in the interruption of protein biosynthesis. However, the most potent and widely demonstrated mechanism involves the cleavage of mRNA by RNase H, a ubiquitous cellular nuclease that degrades the RNA strand of hybrid RNA–DNA duplexes. After the improvements in technology over the last 20 years, one antisense drug (VitraveneTM) has been approved for patients with AIDS-related retinitis,4 and several other molecules are currently in advanced clinical trials. The main problems associated with the use of synthetic oligonucleotides are finding accessible sites on the highly structured mRNA target, their low cellular stability, the difficulty of crossing cell membrane barriers, and the formation of structures that may affect binding affinity and specificity. To overcome these problems, chemists have synthesized oligonucleotide analogues modified almost everywhere in the chain (nucleobases and sugar-phosphate backbone), resulting in molecules with enhanced cellular stability (‘first generation’ of AS-O, e.g. phosphorothioate oligodeoxynucleotides), and in lower toxicity and higher affinity for the target mRNA (‘second generation’ of ASO, e.g. 2′-O-modified oligoribonucleotides). Recently, chemical modifications have provided new DNA and RNA analogues (‘third-generation’ of AS-O), with improved target affinity, biostability and pharmacokinetics, which are expected to generate new antisense-based drugs in the near future.5 As well as the antisense approach, two additional anti-mRNA strategies (Figure 9.1) have been developed, namely ribozymes and RNA interference (RNAi), both

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275

of which also aim to inhibit the expression of a particular gene.6 Synthetic ribozymes mimic the mode of action of naturally occurring ribozymes,7 which bind to mRNA and cleave a precise phosphodiester bond. One of the most commonly studied is the hammerhead ribozyme, which contains two binding arms for substrate recognition and a catalytic domain for cleavage.8 Like AS-O, ribozymes have been chemically modified to increase their stability in cells and their delivery to specific tissues without compromising activity, and some of them are currently in clinical trials for evaluation as therapeutic agents.9 The recent discovery of RNA interference (RNAi)10 has triggered the use of technologies based on this natural gene-silencing mechanism in biological research and their evaluation for the treatment of many diseases.11,12 RNAi is mediated by small interfering RNAs (siRNAs), which are double-stranded oligoribonucleotides typically 21–25 nucleotides in length, and two nucleotide 3′-OH overhangs that are incorporated into an active ribonucleoprotein complex, the RNA-induced silencing complex (RISC).13 Unwinding of the two chains of the siRNA duplex is followed by hybridization of the antisense strand with mRNA, which, depending on the degree of complementarity, either guides RNA cleavage by the activated RISC complex or blocks translation. Synthetic siRNAs seem to be more active than AS-O and could very possibly be developed as a new therapeutic strategy. However, nonspecific effects and toxicity may be associated with high activity, and, although siRNAs are fairly stable in cells, chemical modifications that increase cellular stability and facilitate delivery may be required to ensure their efficiency in vivo for future therapeutic applications.14 Another emerging technology is that based on targeting microRNAs (miRNAs), which are single-stranded nonprotein coding RNAs (21–23 bases in length) that regulate gene expression in plants and animals.15,16 Unlike siRNA-mediated RNAi that inhibits translation of a single gene, an miRNA may target hundreds of different mRNAs. The evidence of miRNA participation in a wide range of cellular functions in mammals (e.g. cell growth and apoptosis, heart and brain development and insulin secretion), together with their involvement in diseases such as neurodegenerative disorders and cancer, makes them attractive new druggable targets.17 Recently, chemically modified anti-miRNA oligonucleotides have proved valuable tools to specifically silence miRNAs in vivo, enabling their functions to be unravelled in a new, promising therapeutic strategy.18,19 Small synthetic oligonucleotides can also be used to target specific sequences of the double helix of DNA (antigene strategy, Figure 9.1).20 These molecules, called triplex-forming oligonucleotides (TFOs),21 recognize and bind polypurine/polypyrimidine tracts in DNA duplexes through non Watson–Crick hydrogen-bonding interactions (Hoogsteen bonding) forming a supramolecular structure that contains three strands. TFOs have been used to selectively inhibit or regulate gene expression at the transcriptional level, and to correct genetic defects arising from point mutations.22 Nevertheless, problems such as the low stability of triple helices, the need to improve their delivery and stability in the intracellular environment and their access to the highly packed nuclear DNA still have to be suitably addressed. In addition to interacting with each other through specific hydrogen bonding, structured nucleic acids (DNA and RNA) also bind to a wide range of target

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molecules, from organic dyes to proteins. Some oligonucleotides (25–40 bases in length), called aptamers, show a highly organized tertiary structure that enables them to bind specifically to their target (generally a protein) with high affinity (Figure 9.1). Aptamers, which can be selected from combinatorial nucleic acid libraries through an iterative in vitro process (SELEX, systematic evolution of ligands by exponential enrichment),23,24 can, in principle, target virtually any protein. Among the many aptamers reported, only MacugenTM, a polyethylene glycol-engrafted oligonucleotide that specifically binds the vascular endothelial grown factor-165 isoform, has been approved for treating age-related macular degeneration.25 Although many aspects such as delivery must be improved to develop new efficient aptamer-based drugs, aptamer technology is emerging as a powerful and versatile tool in various fields of biotechnology, target validation, bioanalysis, diagnostics and imaging.26,27

9.2 Platination as a Tool to Enhance Biological Effects As stated above, one of the requirements for making drugs out of synthetic oligonucleotides is that they efficiently recognize and interact with their targets. Only sufficiently stable complexes can prevent the ribosome machinery from translating the mRNA code into a polypeptide sequence, stop replication or transcription, or block the function of a protein.28 Chemical modification of the chain modulates the affinity of synthetic oligonucleotides for their targets. Substitution of sulfur for one of the oxygens of the phosphodiester bridge (phosphorothioate or S-oligonucleotides),29 which is the modification introduced in the only antisense oligonucleotide approved for therapeutic use,4 slightly decreases the stability of the modified-oligonucleotide–mRNA duplex.30 Conversely, the stability of the duplex can be improved, for instance, by the introduction of a methylene bridge that links the 4′-C and 2′-O and freezes the North conformation of the ribose ring (locked nucleic acids or LNAs),31 or by replacement of the charged sugar-phosphate backbone with noncharged ones, as in peptide nucleic acids (PNAs)32 or morpholino oligonucleotides.33 The stability of the final complex, either duplex or triplex, can also be improved by forming a crosslink between the oligonucleotide and the target. Duplexes can be stabilized by covalently linking their ends. This covalent union has been chemically established by a short oligonucleotide sequence, as in naturally occurring hairpin duplexes, or by nonoligonucleotide linkages (Figure 9.2) such as disulfide bridges,34 alkyl chains,35,36 aromatic moieties such as stilbenes37 or phenanthrenes,38 and metal complexes.39,40 Nevertheless, if oligonucleotides are to be used as therapeutic agents, and recognize and link to their targets, only the synthetic chain can be modified. In that case, a group attached or appended to the synthetic oligonucleotide is expected to give rise to the desired covalent union upon hybridization. Examples of this approach are the use of 2-arylthioethyl- or quinone methide-modified oligonucleotides, as described by Sasaki41 and Rokita,42 respectively (Figures 9.3A and B). In both cases, the target is alkylated upon duplex formation, yielding the interchain crosslink.

Platination to Enhance Biological Effects 277 O

S

N

A N

N

O

O N

B

O

S

N

O

O

(CH2)n

N

N

O

HN

NH

(CH2)3

(CH2)3

NH N

O

O

N

N

O C

NH

O

(CH2)n O

N

N

D O

NH (CH2)n

HN

O

(CH2)n

Ph

N

E

N NH (H2C)3

M

2+

N

= oligonucleotide chain

HN (CH2)3

Figure 9.2 Alternatives to the crosslinking of two oligonucleotide chains by chemical derivatization: disulfide bridges (A), alkyl chains (B), stilbenes (C), phenantrenes (D) and metal complexes (E)

Another alternative that has been pursued is that a metal atom, especially platinum, could be made to establish the crosslink between the synthetic oligonucleotide and the target. In the context of antisense, antigene or anti-microRNA strategies, the platinum(II)-mediated attachment of small synthetic oligonucleotides to their complementary targets (DNA or RNA) is expected to produce a more persistent blockage than that obtained by simple hydrogen-bonding pairing. In this chapter we review the attempts to achieve high therapeutic efficacy as a result of the formation of platinum crosslinks between oligonucleotide chains at the duplex or triplex level. Over the past 15 years, a great deal of effort has been devoted to the development of methodologies to synthesize platinated oligonucleotides, but the scope of applications of such molecules is limited because of the difficulties associated with their preparation (see Sections 9.3 and 9.4).

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Platinated Oligonucleotides

Figure 9.3 Crosslinking promoted by annealing. The reactive species is either a vinyl group (A) or a quinone-methide moiety (B)

Although experiments leading to platinated oligonucleotides have often been followed by crosslinking studies with no clear-cut separation between the two aspects of the study, in this chapter the description of these studies has been split into three sections. Section 9.3 covers most of the methods developed to prepare platinated oligonucleotides, either using preplatinated monomers that are incorporated into oligonucleotide chains, or platinating fully assembled oligonucleotides (partially protected or unprotected, unmodified or containing modified nucleobases). Then, the use of platinated oligonucleotides in studies whose final goal is the formation of crosslinked duplexes, most often to achieve improved antisense effects, is described in Section 9.4. Finally, Section 9.5 reviews the results of experiments aiming at the formation of complexes between platinated oligonucleotides and dsDNA, namely the formation of triple helices (antigene approach). Although the purpose of that study was not to establish a crosslink, it is worth mentioning that the effect of platinating the sense strand of an siRNA duplex was also evaluated in a recent publication.43

Synthesis of Platinated Oligonucleotides

279

9.3 Synthesis of Platinated Oligonucleotides 9.3.1 Platination of Unmodified Oligonucleotides In general, platination of unprotected oligonucleotides in neutral solutions leads to complex mixtures because of the presence of multiple coordination sites at the heterocyclic nucleobases (N7 in guanine, N7 and N1 in adenine, and N3 in cytosine). Although N7 of guanine is the preferred binding site,44 the formation of many different platinated products certainly complicates the isolation of the desired adduct, making its preparation impossible in some cases. An alternative to decreasing the complexity of the crude is to reduce the pH of the reaction medium, in order to keep cytosines and adenines protonated (N3 and N1, respectively) and thus unreactive.45 However, these conditions cause depurination.46,47 Another widely used approach is to reduce the number of guanines to one or two residues, but this alternative prevents the oligonucleotide that is to be platinated from targeting any sequence.48 Given the constraints derived from the structure of platinum(II) complexes together with their preference for guanines, it is possible to control the evolution of the reaction to yield a predominant adduct by choosing the number and position of guanines in the chain as well as the conditions of the reaction. For example, single-stranded oligonucleotides containing two neighbouring guanines react with cisplatin to afford a major 1,2-intrastrand crosslink with platinum bonding both guanines (Figure 9.4A). In contrast, the linear geometry of transplatin prevents formation of these chelates, but it can yield 1,3-intrastrand crosslinks at GNG and ANG sequences (where N is any nucleotide residue other than G) (Figure 9.4B). The geometry of coordination also explains that closure of monofunctional adducts to form interstrand crosslinks is more frequent with transplatin than with cisplatin, especially between the N7 of guanine and the N3 of its base-paired cytosine in a DNA duplex.49 This is because the distortion in the double helix is smaller than with cisplatin-mediated interstrand crosslinks.50,51 Although the preference of transplatin for GNG triplets allows the preparation of platinated oligonucleotides for use in crosslinking experiments (see Section 9.4.2),52 the restriction in sequence imposed by the fact that the oligonucleotide cannot contain other guanines prevents the targeting of any nucleic acid. Additional difficulties associated with this alternative are practical aspects of purification and characterization of GNG-platinated oligonucleotides, and the low stability of the final adduct in some cases. In this respect, it has been reported that the chelates formed by reaction of transplatin with guanines at CGNG sequences rearrange into 1,4-intrastrand crosslinks by platinum migration from the 5′-guanine to cytosine (Figure 9.4C).45,53 The approaches in which transplatin or a binuclear complex reacts with an oligonucleotide to yield a reactive monofunctional adduct are also problematic (see Section 9.4.2).45–48 One difficulty is the low stability of platinated oligonucleotides with one labile bond, since they may react with other nucleobases present in the oligonucleotide (‘suicide’ reaction). In addition, there are many cellular

280

Platinated Oligonucleotides

A

Cl

GG

Pt

Cl

B

Cl H3N

GNG

NH3 NH3

Pt

GG

Here,

H3N

=

Pt

NH3

NH3 Cl

H3N

GNG

Here,

=

CGNG

Here,

=

Pt

N = A, C, T

NH3

C CGNG N = A, C, T

D

Cl

G

H3N

Pt

NH3 PG

Pt

NH3

Cl/H2O

PG H3N Pt

H3N

NH3

G

deprotection

H3N Pt

NH3

G

PG = protecting group = oligonucleotide

Figure 9.4 Reactions of cisplatin with GG-containing oligonucleotides (A) and of transplatin with GNG-containing chains (N = A, C, T) (B). trans-Pt(NH3)2 1,3-chelates isomerize to 1,4chelates when the GNG sequence is flanked by a 5′ C (C). The reaction of guanine-containing oligonucleotides with suitably protected platinum complexes affords platinated oligonucleotides with one labile bond (D)

nucleophiles that may render the complex unreactive before it reaches its target. One solution envisages transplatin being replaced by a trans-(NH3)2 platinum complex in which the metal is linked to one labile substituent and a protecting group selectively removable upon interaction with the target. However, this ideal protecting group has not yet been discovered. So far, only thymine derivatives (Pt-N3 linkage) have been used as protecting groups in the preparation of platinated oligonucleotides with one labile bond. Their use has not been extended, however, because they are labile to acidic conditions that may induce depurination (Figure 9.4D).46,54 Over the years, chemists have applied various methods to clarify the structure of platinated oligonucleotides. The presence of platinum had been assessed by atomic absorption or induced coupled plasma mass spectrometry, but these techniques were replaced by modern mass spectrometric techniques that allow the ionization of oligonucleotides, such as ESI or MALDI (ESI = electrospray ionization, MALDI = matrix-assisted laser desorption ionization). The formation of a coordination compound was inferred from the evidence that reaction of the platinated oligonucleotide with CN− or thiourea had afforded the nonmodified chain. NMR

Synthesis of Platinated Oligonucleotides

281

provides both proof of the presence of platinum (when the use of 195Pt enriched complexes allows 195Pt spectra to be recorded) and, based on the comparison between the chemical shifts of platinated and nonplatinated oligonucleotides, information on the coordination point. This item of information, however, has also been obtained from gel electrophoresis analysis after chemical reactions that modify nonplatinated nucleobases and cleave the chain, and, more recently, from mass spectrometric analysis after digestion with exonucleases. Both NMR and X-ray crystallography provide structural information at the atomic level. The first structure was solved more than 10 years ago, and showed the distortion caused in a duplex by the formation of a 1,2-intrachain cis-[Pt(NH3)2] adduct at a GG doublet.55 Highresolution pictures have also been obtained for duplexes with one strand coordinated to other platinum complexes,56,57 as well as for cis-58 and trans-[Pt(NH3)2] crosslinked duplexes.51 9.3.2 Platination of Unprotected Oligonucleotides with an Appending Chelating Group The McLaughlin group explored the alternative of platinating oligonucleotides incorporating an ethylenediamine group.59,60 They introduced the phosphoramidite derivative of the bis-Fmoc-protected chelator (Fmoc = 9-fluorenylmethoxycarbonyl) at the 5′-end of an immobilized oligothymidine. After ammonia treatment and purification of the oligonucleotide derivative, reaction with K2PtCl4 afforded the expected dichloroplatinum(II) derivative. However, even though formation of a chelate favours coordination with the diamine, the high reactivity of K2PtCl4 precludes the inclusion of nucleobases other than thymine and thus limits the scope of this approach (Figure 9.5).

Fmoc HN

NiPr2 N P OCNE Fmoc HO Tn

H2N

NH

Tn

K2PtCl4

NH

H2N Cl

Pt

Tn

Cl

= solid matrix Fmoc = 9-fluorenylmethoxycarbonyl CNE = 2-cyanoethyl

Figure 9.5 Post-assembly platination of an oligonucleotide bearing an appending chelator

9.3.3 Solid-Phase Synthesis Using Platinated Building Blocks Biopolymers, such as peptides and oligonucleotides, which are made up of the repetition of a basic unit (amino acid or nucleoside, respectively), are commonly assembled through solid-phase technologies. This process can be automated because the

282

Platinated Oligonucleotides

incorporation of each synthon always involves the same series of steps, namely coupling of a suitably protected building block and removal of a temporary protecting group to liberate the functional group to which the next unit has to be attached. The elongation of oligonucleotide chains in automatic synthesizers utilizes the phosphite triester approach,61 which is the most effective methodology, even though it involves the largest number of steps. Nevertheless, the H-phosphonate method62,63 has also provided satisfactory results for the preparation of some oligonucleotide analogues. It can be used in automatic synthesizers, provided that the standard synthesis cycles are suitably modified. The preparation of platinated oligonucleotides from platinated nucleoside monomers was first described in 1996 by two separate groups.64,65 Both aimed at introducing platinum at an internal position of the oligonucleotide chain, and used the platinated H-phosphonate nucleoside derivatives depicted in Figure 9.6A, which shows the differences in the nucleobase involved, as well as in the nature of the ligands and the stereochemistry of the complexes. Cech and coworkers64 argued that H-phosphonates were more robust than phosphoramidites, and that phosphitylation had to precede platination.

NH3

O

A

N DMT O

O

N

Pt O

Bhoc

H3N

O

N

O H P O

O

O

NH N

NH2

Cl NH3

N

NH Fmoc

O

N

NH3 DMT O

Cl

Pt

O H P O

H3N Pt O N HN

B

H3N Cl

O NH

N

O OH

DMT = 4,4'-dimethoxytrityl Bhoc = benzhydryloxycarbonyl Fmoc = 9-fluorenylmethoxycarbonyl

Figure 9.6 Structures of the 2′-deoxythymidine- and 2′-deoxyguanosine H-phosphonate monomers used to assemble platinated oligonucleotides by stepwise solid-phase synthesis (A), and of the platinated PNA monomer used to obtain platinated PNAs (B)

Synthesis of Platinated Oligonucleotides

283

H-phosphonate chemistry was used throughout chain elongation. They found that it was more difficult to incorporate the platinated synthon than the natural nucleoside H-phosphonates. Final deprotection with concentrated aqueous ammonia afforded crude oligonucleotides showing rather complex HPLC profiles, and the coordination sphere of the metal was modified. Pyridine had replaced chloride, yielding platinated oligonucleotides with no potential to react with the nucleophiles of a complementary chain. Side reactions during final deprotection and work-up were held to account for the low quality of the crudes. The same sequence of steps was described in Lippard’s study,65 but Hphosphonate chemistry was only used to introduce the platinated monomer, while standard phosphite triester chemistry was used for all the other nucleosides. The authors indicated that after incorporation of the platinated unit the synthesis cycle was modified, but no details were given. Elongation of the full oligonucleotide chain was followed by incubation in water to allow a 1,2-intrastrand G–Pt–G chelate to form, and then by the cleavage and deprotection treatment. A fairly pure platinated oligonucleotide, albeit in low yield, was obtained, but the authors indicated that the protocol still required optimization. All these results indicate that the introduction of platinated nucleoside building blocks within the chain was neither straightforward nor easy to optimize. Although labile substituents seemed to be compatible with oligonucleotide elongation using phosphoramidites,65 subsequent studies from other groups showed that they could react with phosphoramidite derivatives.60,66 The attempts of the groups of van Boom, Reedijk and Lippert to prepare platinated oligonucleotide analogues, in particular peptide nucleic acid chains (PNAs), by solid-phase synthesis are also worth mentioning here.67 The N-Fmoc-N2-Bhoc (Bhoc = benzhydryloxycarbonyl) guanine-containing PNA monomer was reacted with a transplatin analogue to afford an N7-platinated guanine building block with a labile chloride ligand (Figure 9.6B). This monomer was then introduced at the N-terminal of resin-linked, protected PNA chains. Then, following treatment with trifluoroacetic acid, which removes all protecting groups, target-platinated PNAs were obtained. The key point for the success of the synthesis is that the final treatment is applied in conditions that do not block the reactive position at the metal centre. This contrasts with the effect of the ammonia treatment used for the final deprotection of oligonucleotides, which provides an unreactive platinated oligonucleotide.

9.3.4 Regioselective Platination of Partially Protected Oligonucleotides: Use of O6-Guanine Protecting Groups The van Boom and Reedijk groups explored an elegant alternative, in which platinum is directed to the desired guanines while the reaction with other guanines is prevented by the introduction of a bulky protecting group at the O6 position.68,69 The preliminary study68 found that N2-protected guanines could be platinated at the N7 position, unlike O6,N2-protected ones that could not (Figure 9.7). It also

284

Platinated Oligonucleotides Pt

O N O

O

NH

N

N

NH PG

Platinum(II) complex (at least one labile bond)

O

O

N O

O O

N

NH N

NH PG PG = protecting group

N O

O

N O

O O

N

N N

Platinum(II) complex

NH PG

(at least one labile bond)

PG = protecting group

Figure 9.7 N2-protected guanines react with platinum complexes, but protection of the O6 with a bulky group prevents platination at the N7 position

showed that fully protected oligonucleotides reacted very slowly with platinum(II) complexes. The yield of the platination reaction increased when the phosphate protecting groups were removed, which was associated with favourable electrostatic interactions between the positively charged cation and negatively charged phosphates. The proof of concept was obtained when different 2′-deoxyhexanucleotides were selectively platinated at the guanines whose O6-position had been left unprotected (Figure 9.7).69 The oligonucleotide chains were assembled on controlled pore glass (CPG) using an oxalyl linker and either the standard phosphoramidite derivatives (T, ABz, CBz, GiBu, where Bz = benzoyl, iBu = isobutiryl) or the N2-propanoyl,O6diphenylcarbamoyl phosphoramidite derivative of guanine. Cleavage and removal of phosphate-protecting groups afforded the desired partially protected oligonucleotide, which reacted regioselectively with [Pt(en)Cl2] (en = 1,2-diaminoethane). The target platinated oligonucleotide was obtained after removal of nucleobase protecting groups with concentrated aqueous ammonia. Although the authors suggested that this approach might be used for the generation of other site-specifically platinated oligonucleotides, such as trans-Pt-1,3-GNG adducts, this alternative has not yet been explored. Moreover, this alternative seems to be restricted to the preparation of platinated oligonucleotides with no labile platinum–ligand bonds, unless a platinum protecting group is made available that is stable to ammonia, but removable under conditions that do not damage the oligonucleotide.

Synthesis of Platinated Oligonucleotides

285

9.3.5 Attachment of Platinated Units to Protected Oligonucleotides A different approach for the preparation of platinated oligonucleotides is the incorporation of the platinated unit to the growing oligonucleotide chain instead of using a platinated nucleoside derivative. In 1999, Ren, Cai and Segal studied the reaction of a [Pt(en)Cl2] analogue containing an appending hydroxyl group with either a phosphitylating reagent or a phosphoramidite derivative of thymine in the presence of tetrazole (or a tetrazole derivative).66 In the two cases the hydroxyl group reacted with the phosphorus(III) derivative (Figure 9.8A), but the evolution of the resulting complex was different depending on whether the reaction yielded a phosphoramidite or a phosphite triester. In the first case, the nucleophilicity of phosphorus prompted quick displacement of one of the chlorides linked to platinum, affording a double chelate. This platinated phosphoramidite could not be activated by tetrazole. Therefore, it did not react with free hydroxyl groups in a nucleoside or in an oligonucleotide chain. The phosphite triester also behaved as a nucleophile, but the resulting complex was a mixture containing a double chelate and the target [Pt(en)Cl2]-derivatized nucleotide. The same group also described70 the synthesis of platinum complexes with no chelating amines, also containing hydroxyl groups to allow attachment to oligonucleotides (Figure 9.8B). However, to the best of our knowledge this methodology has not been explored so far. NiPr2

NiPr2

A CNEO

P

NiPr2

tetrazole

NH

H2N

Pt Cl

Cl iPr2N CNEO

Cl

OH (CH2)n Pt

NH

R-OH

Pt

OCNE

Cl P O tetrazole CNEO NiPr2

Cl

OCNE

tetrazole P O

O

T

O DMT

Cl

H2N

Cl

H2N

NH Pt

Cl

B

O

P

OH

Pt Cl

NH

H2N

Cl

O P O CNEO

O

T

O DMT

P

O

NH Pt

Cl

O

NH2

O

T

O DMT

Cl

CNE = 2-cyanoethyl DMT = 4,4'-dimethoxytrityl

NH2 NH2 (CH2)n OH

Figure 9.8 Intramolecular reaction of phosphorus(III) reagents [(2-cyanoethoxy)bis(diisopropylamino)]phosphine and 5′-phosphoramidite derivative of 3′-O-DMT-thymidine] with derivatized [Pt(en)Cl2] complexes (A). Structure of a cisplatin analogue with two appending hydroxyl groups (B)

286

Platinated Oligonucleotides NH2Me

O N N

Pt O

NH2

(CH2)6

OH

NH2Me activator iPr2N P O TTTT

protecting group

[O]

NH3

NH2Me

aq HCl pH 2,3

OCNE

Cl

Pt

O NH2

(CH2)6 O P O TTTT

NH2Me

O

= solid matrix

CNE = 2-cyanoethyl [O] = oxidation (phosphite to phosphate)

Figure 9.9 Platinated oligothymidines with one labile bond are obtained by reacting the phosphitylated, resin-linked oligothymidine and a masked platinum complex with an appending hydroxyl group, followed by removal of the protecting groups

At the same time, the van Boom, Lippert and Reedijk groups developed a parallel alternative to tether a platinum complex with one labile bond to oligonucleotides.54 Their strategy involved phosphitylation of the 5′-hydroxyl of a resinlinked oligonucleotide chain, and the subsequent reaction of the resulting phosphoramidite with a platinum complex that contained a hydroxyl group (Figure 9.9). After the oligothymidine deprotection step with aqueous ammonia, treatment with aqueous HCl (pH = 2.3) removed the platinum protecting group and introduced a labile chloride group, as required for crosslinking to a complementary sequence. Further development of this approach, in particular optimization of the platinum protecting group to render its elimination compatible with any oligonucleotide sequence (in other words, with no risk of depurination), has not yet been reported. 9.3.6 Regioselective Platination of Base-Modified Oligonucleotides Platinum(II) complexes can react with the nitrogen and sulfur nucleophiles of amino acid side chains, as well as the heterocyclic nitrogen of nucleobases. Since the formation of Pt–S bonds is the kinetically favoured reaction and many sulfur nucleophiles are present in the cells, several research groups have studied the possible transfer of platinum from sulfur to nitrogen and the competition between both nucleophiles.71–81 These studies showed the preference of platinum(II) complexes for sulfur-containing species. Then, depending on the structures of the substrate and the platinum(II) complex (stereochemistry, number of labile bonds), the initially formed Pt–S adduct can evolve into monofunctional, bifunctional and even trifunctional adducts in which Pt–S bonds are usually replaced by the thermodynamically more stable Pt–N linkages. The reaction between several peptide–oligonucleotide conjugates and platinum(II) complexes such as cisplatin, Pt(dien) (dien=1,5-diamino-3-azapentane), Pt(en) and transplatin, revealed that some S–Pt–N adducts, in which the metal linked methionine and either histidine or guanine, were quite stable. In

Synthesis of Platinated Oligonucleotides N S

XS

N O

N XI

H3N

HN

HN N

N

NH3 S Pt

NH

HN

Cl

O

287

oligonucleotide (A, C, G, T)

H3N

Pt

NH3 Cl

N N H

NH

N N

N O

N

O

oligonucleotide (A, C, G, T)

Figure 9.10 Modified oligonucleotides containing the four nucleobases and two cytosine analogues can be selectively platinated at the appending thioether and imidazole tethers. (Adapted from J. Biol. Inorg. Chem., 2007, 12, 901–911.)

addition, platinum preferred the N7 of guanine rather than nitrogens from the imidazole ring. However, thioether–platinum–imidazole adducts could form in the presence of guanines, provided sulfur promotes binding to the imidazole ring rather than the N7 of guanines.78,80 On the basis of these studies, Marchán and Grandas82 envisaged an alternative method for preparing modified oligonucleotides that included, in addition to all the natural nucleobases, the thioether and imidazole functional groups present in the methionine and histidine side chains. Sulfur was expected to direct the platination process and prevent uncontrolled reaction with the nucleobases (Figure 9.10). For this purpose, two 5-methylcytidine analogues with thioether or imidazole groups attached to the 4-position (XS and XI, respectively; see Figure 9.10) were synthesized by reacting protected 4-(1,2,4-triazole)cytidine with the corresponding amines.82 After phosphitylation, the phosphoramidite derivatives were successfully introduced in oligonucleotide chains using the standard phosphite triester chemistry with no protection on either the thioether or the imidazole ring. Several oligonucleotide sequences containing either one or two analogues appended to the 5′-end were synthesized and used in preliminary platination studies. The modified nucleosides were placed at the 5′-end to minimize the destabilization that modified cytosines cause in a DNA duplex, especially in the middle of the chain.82,83 The platination studies were carried out using transplatin, because it is a simple, mononuclear complex whose reactions with single-stranded oligonucleotides have been thoroughly studied, and whose capacity to form crosslinked duplexes is clearly established.49–51 Unlike cisplatin, transplatin offers the advantage that the trans geometry prevents the sulfur ligand from labilizing the trans ammine group. Hence, using transplatin, mixtures of di- and tricoordinate adducts, which are more difficult to handle, are not formed.80 The reaction between transplatin and oligonucleotides containing the thioethermodified nucleobase was fast, but generally afforded unstable adducts and complex reaction mixtures. However, the imidazole-containing oligonucleotides reacted with transplatin much more slowly, in particular in slightly basic solutions, and it was found that imidazole-modified cytosine was less reactive than the natural nucleobases.

288

Platinated Oligonucleotides

Figure 9.11 Identification of the platination site in modified oligonucleotides. Use of chemical and enzymatic reactions in combination with mass spectrometric data. (Adapted from J. Biol. Inorg. Chem., 2007, 12, 901–911.)

In contrast, platination of oligonucleotides containing the four natural nucleobases and the two nucleoside analogues XS and XI in neighbouring positions (5′dXXACGTTGAG) afforded fairly stable chelates, in which the trans-Pt(NH3)2 unit linked the thioether and the imidazole ring. The new compound isolated from the reaction of 5′dXSXIACGTTGAG with transplatin had the mass expected for a platinated oligonucleotide. It did not react with H2O2, which indicated coordination to the thioether. It was more stable to digestion with a 5′-exonuclease (calf spleen phosphodiesterase) than the nonplatinated oligonucleotide, which also confirmed coordination to the thioether at the 5′-end. Finally, digestion with a 3′-exonuclease (snake venom phosphodiesterase) removed the natural nucleosides, giving the 5′ dXSXI fragment plus a trans-Pt(NH3)2 unit, confirming coordination with the imidazole-containing nucleoside (Figure 9.11). Four chelates may be formed, because platinum may react with either of the two sulfur nonbonding electron pairs and link to either of the two nitrogens of the imidazole ring. The presence of a single main peak in the HPLC traces indicated that the isomers coeluted in the analysis conditions used. Similar results were obtained with 5′dXIXSACGTTGAG. The two chelates involving XI- and XS-platinum bonds remained unmodified for 24 h in the reaction mixture. However, at longer reaction times platinum migrated from the thioether to the N7 of the closest guanine in the sequence, to give a chelate that was thermodynamically more stable. In no case were monofunctional adducts detected, which indicates that the reaction of platinum with the thioether was immediately followed by coordination with one of the nitrogen atoms in the imidazole ring. Moreover, although the two modified oligonucleotides contained the four nucleobases and a GAG triplet, transplatin (in equimolar amount in relation with the modified oligonucleotide) preferred the thioether and the imidazole groups of the two neighbouring cytosine analogues to any of the natural bases. In other words, these modified, but unprotected, oligonucleotides were regioselectively platinated in spite of the presence of several guanines in the chain. These results opened up the possibility of using these chelates in crosslinking experiments, aiming at the control of gene expression by targeting complementary sequences in either mRNA or miRNAs.84

Platinated Oligonucleotides for Duplex Crosslinking

N

Pt

N

HN

N N

H2 N

NH2

N

O

NH2

H2N

NH3

NH Cl

N N

O

Cl

Pt

NH3 NH3

289

NH3 NH3

N N

O

= oligonucleotide (protected when anchored to resin) = solid matrix

Figure 9.12 Introduction of a chelating group in oligonucleotides using a convertible nucleoside, and regioselective reaction of the polypyrimidine chain with cisplatin.

While studying the effect of platinated aminoalkyl arms on triplex formation, Miller et al. reported the synthesis of platinated oligopyrimidines in which one cytosine was replaced by N4-(2-aminoethyl)cytosine.85 The oligonucleotides were assembled on a solid support, and the incorporation of a 4-triazole-deoxyuridine derivative in the middle of the chain allowed the introduction of the aminoethyl function by treatment with ethylenediamine. After ammonia deprotection and cleavage, the aminoethyl arm of the unprotected oligonucleotide was transformed into a Pt(dien) or cis-chlorodiammineplatinum(II) complex by reaction with [Pt(dien)Cl]Cl or cisplatin, respectively (Figure 9.12).

9.4 Use of Platinated Oligonucleotides for Duplex Crosslinking 9.4.1 Initial Experiments In 1983, Vlasov et al. described for the first time the use of platinum(II) compounds for crosslinking DNA duplexes.86 The first step was the reaction of a binuclear platinum(II) compound incorporating two labile functions with the N7 of an oligonucleotide containing a single guanine. Then, hybridization promoted reaction with the guanine at the 5′ end of the complementary strand and gave the interstrand crosslink (Figure 9.13). Regioselectivity in the platination of the second strand, together with the fact that noncomplementary chains were not crosslinked, showed that complementarity is a key point in these processes. As a result, the authors envisaged the possibility of using platinated oligonucleotides for damaging certain sequences of nucleic acids in vivo. Given the great affinity of phosphorothioate groups for platinum(II) complexes,87 a few years later Orgel et al. introduced a single phosphorothioate group at the 5′-end of an antisense oligonucleotide to mediate crosslinking to longer DNA strands that contained a complementary sequence,88,89 or even to proteins.90 In these

290

Platinated Oligonucleotides 5'

dTCCGCCTTT + Pt Pt

Pt

3'

5'

dTCCGCCTTT Pt

pH 5.0

Pt

dAGGCGG

5'

dTCCGCCTTT Pt

pH 7.1

Pt

3'

dAGGCGG

Pt = binuclear platinum(II) complex

Figure 9.13 Regioselective reaction with a binuclear complex is followed by hybridization to the complementary chain, which triggers formation of the crosslink

3' 3'

5'

S Pt

5'

Figure 9.14 Oligonucleotides with one phosphorothioate group are regioselectively platinated at sulfur. Then duplex formation induces formation of an interchain crosslink. (Adapted from Nucleic Acids Res., 1990, 18, 5163–5171.)

experiments, the platinum(II) compound (cisplatin, K2PtCl4, transplatin or a binuclear platinum complex) was added once hybridization of both strands was achieved, as demonstrated by nondenaturing gel electrophoresis. Then, crosslinking occurred when the single-stranded tail of the template bent around the 5′-terminal phosphorothioate of the antisense oligonucleotide, which placed sulfur and residues 2–5 in the target strand sufficiently close to be crosslinked by the platinum(II) compound (Figure 9.14). The different assays showed that guanine and adenine and, to a lesser extent, cytosine were the preferred sites for crosslinking, but, in the absence of these residues in positions 2–5, crosslinking occurred at thymine. The antisense oligonucleotides contained few guanines and when a guanine was placed close to the phosphorothioate group, an intramolecular ‘suicide’ reaction occurred. Among the platinum(II) compounds, the crosslinking efficiency of transplatin was comparable to that of K2PtCl4, but cisplatin was less efficient. 9.4.2 Crosslinking Experiments Using Platinated Natural Oligonucleotides and PNA Analogues While conducting research into the interactions between cis- and transplatin and DNA, Leng et al. discovered a linkage isomerization reaction between transplatinmodified oligonucleotides and their complementary strands, promoted by the formation of a DNA double helix.52 Although oligonucleotides with trans-Pt(NH3)2 1,3-intrastrand crosslinks at GNG triplets (where N is any nucleoside but G) were kinetically inert as long as they remained single-stranded (except those containing

Platinated Oligonucleotides for Duplex Crosslinking Cl H3N

GNG

Pt

NH3 Cl

GNG

N = A, C, T

+ complementary chain, hybridization

GNG

GNG

C N' C

C N' C

= oligonucleotide

291

,

H3N

=

N': base complementary to N Pt

NH3

Figure 9.15 Hybridization of an oligonucleotide with a 1,3-trans-Pt(NH3)2 chelate to the complementary strand induces a rearrangement that crosslinks the two chains

the CGNG sequence, see Section 9.3.1 and Figure 9.4C),45,53 they became unstable upon pairing with their complementary DNA or RNA strands. This interaction promoted the rearrangement of the 1,3-intrastrand chelate into an interstrand crosslink between the 5′-G of the initially platinated triplet and its complementary C, resulting in the covalent linkage of both strands (Figure 9.15). Since isomerization kinetics were independent of the nature and concentration of salts in the reaction medium, it was suggested that the mechanism involved direct nucleophilic attack of the cytosine complementary to the platinated 5′-G residue. The process was not dramatically affected by the nature of the residue (A, C or T) lying between the chelated guanines. Since the discovery of this reaction, some authors have attempted to optimize the isomerization process and make platinated oligonucleotides efficient therapeutic molecules for the control of gene expression.91 The rate of the interstrand crosslinking reaction varied, depending on the sequence facing the intrastrand crosslink, and was too slow when the complementary oligoribonucleotide sequences were 5′-CN′C (N′ = any nucleotide). However, by using platinated 2′-Omethyloligoribonucleotides and modifying the complementary sequence so that the triplet facing the chelate was replaced by 5′UA or 5′CA doublets, the rearrangement was achieved within a few minutes.92 In an experiment with a 12/11-mer in which a d(AT) doublet replaced the CTC triplet in a regular duplex, an interstrand G-N7– Pt–A-N1 crosslink was formed.93 Although this methodology has given promising results in blocking translation both in vitro and in cultured cells,94,95 its application in therapy is problematic. Again, sequence restriction requires a maximum of two guanines in the GNG triplet in the oligonucleotide to be platinated. The same strategy was used to crosslink an antisense oligo-2′-Omethylribonucleotide to the human telomeric sequence (T2AG3)n.96

292

Platinated Oligonucleotides

A different approach to crosslink platinated oligonucleotides with their complementary strands was developed by Lippert et al.97 It was based on generating a transplatin monofunctional adduct in a single-stranded oligonucleotide, which, upon hybridization with the complementary chain, evolved into an interstrand crosslink. To minimize the formation of multiplatinated side products, the reaction was carried out at low pH (3.6) and temperature (4 °C), and the oligonucleotide to be platinated contained a single guanine at the 5′-end. The corresponding N7-Pt(NH3)2Cl adduct reacted with the opposite guanine in the complementary strand to form the final interstrand crosslink. Broad application of this strategy is severely limited by the above-stated requirements and by the fact that monofunctional adducts may be easily deactivated by intracellular nucleophiles before reaching the therapeutic target. As also stated above (Sections 9.3.1 and 9.3.5), use of a platinum protecting group might at least prevent undesired replacement of labile ligands in monofunctional adducts.46 As in the previous strategy, formation of an interstrand crosslink between a PNA and the complementary DNA strand was achieved by introducing a transplatin monofunctional adduct at a guanine of the PNA chain.67 Unlike the previous approaches, in this case there was no sequence restriction because the platinated PNA was synthesized by standard solid-phase methodologies that used a platinated monomer of guanine (see Figure 9.6B). However, the crosslinking reaction was too slow, which was attributed to the low solubility of the platinated PNA in the reaction media.

9.4.3 Crosslinking Experiments Using Oligonucleotides Platinated at Base-Modified Nucleosides As oligonucleotides regioselectively platinated at thioether and imidazole groups were made available,82 studying whether hybridization with the complementary chain would promote platinum migration and generate an interstrand crosslink was the next step. Annealing of platinated oligonucleotide 5′dXIXSACGTTGAG (coordination positions are highlighted in bold) to an equimolar amount of the complementary chain (5′dCTCAACGTGTTTG) triggered a ligand-exchange substitution, affording exclusively an interchain crosslink product, as detected by both anion exchange HPLC and polyacrylamide gel electrophoresis (Figure 9.16).84 The crosslinked duplex was isolated and its structure was inferred from MALDI-TOF mass spectrometric analysis before and after chemical and enzymatic reactions and from NMR. It was confirmed that platinum remained coordinated to imidazole in the modified nucleoside XI, but linked to a new position, the N7 of the guanine opposite XS. Similar results were obtained using platinated 5′dXSXIACGTTGAG. Although the crosslinking reactions were slow, under the hybridization-favouring reaction conditions and in the presence of the complementary chain, the chelates (5′dXIXSACGTTGAG or 5′dXSXIACGTTGAG) remained stable. In other words, under these conditions the intrachain rearrangement to a more stable chelate (5′dXIXSACGTTGAG or 5′dXSXIACGTTGAG) was not observed.

Platinated Oligonucleotides for Duplex Crosslinking

293

Figure 9.16 Duplex formation between oligonucleotides selectively platinated at nucleobases modified with appending thioether and imidazole groups (XS and XI, respectively) and the complementary chain promotes migration of platinum from sulfur to guanine (N7), and crosslinks the two chains. (Adapted from Angew. Chem. Int. Ed., 2006, 45, 8194–8197.)

Figure 9.17 Crosslinked duplexes can be obtained by reacting transplatin, an oligonucleotide incorporating the XS and XI nucleobases at neighbouring positions and the complementary chain. The metal links the imidazole ring and the N7 of the guanine opposite XS. (Adapted from Angew. Chem. Int. Ed., 2006, 45, 8194–8197.)

These results showed that this methodology82,84 allowed targeting and crosslinking of complementary oligonucleotides without sequence restriction, and they opened up the possibility of using such platinated oligonucleotides in experiments aiming at the control of gene expression, targeting either mRNAs or miRNAs. The possibility of obtaining crosslinked duplexes without previous platination of one of the two strands was also explored, as Orgel and collaborators had done earlier.88,89 One-pot crosslinking experiments were carried out by mixing equimolar amounts of the modified oligonucleotide, the complementary chain and transplatin in duplex-forming conditions (Figure 9.17). The crosslinked duplex formed was the same as in the previous experiments, which indicated that previous isolation of the platinated modified oligonucleotide was unnecessary. Moreover, the thioether–Pt– imidazole chelate was always detected in the one-pot reaction mixture, which suggested that its formation preceded that of the crosslinked duplex. The fact that no

294

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other chelates were detected again confirmed that the high affinity of sulfur for the metal led to the formation of the intrachain thioether–Pt–imidazole adduct, preventing other oligonucleotide sequences from being platinated, even when they contained various guanine residues. The ligand-exchange substitution process was highly regioselective, since, in crosslinking experiments involving platinated 5′dXIXSACGTTGAG and complementary oligonucleotides differing in the overhang sequence, migration from the thioether to the guanine opposite XS was always preferred, even over migration to the guanine opposite XI. Noncomplementary strands did not become crosslinked, which confirmed that crosslinking was the result of annealing-promoted rearrangement. Hence, the crosslinking process was also sequence selective, since it only took place if the two chains were complementary. Although reaction times were high, particularly when the modified oligonucleotide was not preplatinated, changes in the modified oligonucleotide and the complementary chain could accelerate the crosslinking reaction, as shown by Giraud-Paris and Leng in their experiments with oligonucleotides containing a platinated GNG triplet.92,94

9.5 Use of Platinated Oligonucleotides for Triplex Crosslinking As previously stated (Section 9.1), one of the problems of antigene strategy is the low stability of triple helices in physiological conditions, as the third strand can be displaced by the replication or the transcription machineries. Taking into account the requirements of the parallel motif for triplex formation (polypyrimidine oligonucleotides recognize polypurine-polypyrimidine tracks of DNA), in 1996 Leng and coworkers attempted to covalently attach the third oligonucleotide strand to the duplex by using transplatin as the crosslinking reagent.98 Then, polypyrimidine oligonucleotides containing a single guanine either in the middle or at the 5′-end of the sequence were platinated at acidic pH, which enabled the regioselective formation of the monofunctional adduct at the N7 of guanine. Both platinated oligonucleotides bound the ‘complementary’ duplex and formed interstrand crosslinks that involved almost exclusively guanines within the purine-rich strand. The crosslinking yield was higher when the monofunctional adduct was located at the 5′-end of the third strand because the triplex was more stable than when the platinum adduct was located in the middle of the sequence. In addition, several oligonucleotides containing a monofunctional adduct at cytosine were evaluated in crosslinking experiments and, although the triplex partially dissociated, the interstrand crosslinks were also formed. The rate of the crosslinking reaction depended on the nature of the adduct, but not strongly on its location.99 The problem associated with this strategy is, again, the low stability of monofunctional adducts, which may react with many different compounds (glutathione, proteins) before reaching dsDNA. Moreover, as reported by Leng et al.,98 platinated oligonucleotides with one labile bond can be deactivated by suicide reactions to form intrastrand crosslinks. More recently, Miller et al. examined the effect of oligonucleotides platinated at N4-(2-aminoethyl)cytosines85 (with Pt(dien) or cis-Pt(NH3)2Cl moieties) in

References

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triplex stability (see Figure 9.12). As in the results reported previously,59,60 the Pt(dien) group sterically interfered with the formation of the triplex when it was located in the middle of the sequence, but triplex stability was restored when this group was tethered to the 5′-end of the triplex-forming oligonucleotide via a 2-aminoethylcarbamate linkage. In crosslinking experiments, the cisplatin monoadduct-containing oligonucleotide did not produce any significant interstrand crosslink with the DNA duplex, which was again attributed to steric hindrance. As indicated in Section 9.3.2, the McLaughlin group had reported the synthesis of oligothymidine sequences tethering a cis-bifunctional platinated complex at the 5′-end, for use in DNA duplex targeting by oligodeoxynucleotide-mediated triplex formation.59,60 These platinated oligonucleotides retained crosslinking ability because of the presence of the two cis chloride ligands attached to the platinum centre. Although a dT8 sequence was not long enough to facilitate triplex formation and hence platinum-mediated crosslinking, the dT15 sequence formed a triplex and crosslinked duplexes containing two noncomplementary overhangs and a GpG sequence at various sites along the overhang. Platinum-mediated crosslinking occurred exclusively with the purine-rich strand of the duplex target directly involved in triplex formation, and replacement of guanines by thymines or 7-deazaguanines in this strand prevented the formation of the interstrand crosslink. Unfortunately, no further progress has been reported.

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96. Perrier, S.; Seela, F.; Schwartz, A.; Leng, M.; Chottard, J.C.; The human telomeric sequence (T2AG3)n is efficiently cross-linked by AN1 binding to the platinum of a trans-Pt(NH3)2 chelate of an antisense oligo-2′-O-methylribonucleotide; Eur J. Inorg. Chem., 2003, 8, 1641–1644. 97. Muller, J.; Drumm, M.; Boudvillain, M.; Leng, M.; Sletten, E.; Lippert, B.; Parallel-stranded DNA with Hoogsteen base pairing stabilized by a trans-[Pt(NH3)2]2+ cross-link: characterization and conversion into a homodimer and a triplex; J. Biol. Inorg. Chem.; 2000, 5, 603–611. 98. Colombier, C.; Lippert, B.; Leng, M.; Interstrand cross-linking reaction in triplexes containing a monofunctional transplatin-adduct; Nucleic Acids Res., 1996, 24, 4519–4524. 99. Bernal-Mendez, E.; Sun, J.; Gonzalez-Vilchez, F.; Leng, M.; Reactivity of transplatinmodified oligonucleotides in triple-helical DNA complexes. New J. Chem., 1998, 22, 1479–1483.

10 Rhodium– and Tin–DNA Interactions and Applications Kenneth D. Camm and Patrick C. McGowan

10.1 Introduction The ubiquity of transition metals employed in biological processes is a direct consequence of the versatile chemistry such metals exhibit. In particular, the multiple oxidation states and diverse structural motifs accessible to transition metals have resulted in their incorporation into complex ‘bioinorganic’ species in nature. These same properties have facilitated the development of artificial transition metal complexes, designed to modify or regulate many biological processes.1 The Lewis acidity of transition metals, and consequently their affinity for Lewis bases such as N, O or S donor ligands, renders transition metal complexes suitable for interaction with a multitude of biologically relevant species, including nucleic acids (DNA and RNA). Cellular function and replication relies on the chemistry and interactions of nucleic acids, and consequently these species have frequently been targeted in therapeutic protocols for diseases as varied as cancer and arthritis.2 The first, and to-date most successful, inorganic complex employed in the treatment of cancer, [Pt(NH3)2Cl2] (cisplatin, Figure 10.1a)3,4 and its derivatives, is highly effective against testicular and ovarian cancer, and in other malignancies including cervical, bladder, head and neck tumours. The cytotoxic activity of cisplatin is a direct consequence of covalent interactions between the Pt centre and N7s of adjacent guanine residues on DNA, to form a covalent 1,2-intrastrand d(GpG)crosslink.4 Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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Crosslinking results in inhibition of transcription and DNA replication, eventually leading to cell death. Following the success of cisplatin, a catalogue of research has accumulated, investigating the interactions of other transition metals with DNA.1,2,5–9 Much research has focused on the development of novel therapeutic protocols that may circumvent the toxicity issues associated with cisplatin chemotherapy, and to probe the nature of nucleic acid structure and function in biological processes. To this end, investigations have been performed on metal–DNA interactions covering a large majority of the periodic table, many of which are reviewed in this publication. Herein we describe recent developments focusing on the interactions of rhodium and tin with DNA, and how such work has contributed to our understanding of the role of nucleic acids in biological processes and facilitated the development of novel antitumour agents.

10.2 Metal–DNA Interactions There are three principle interactions that occur between metal complexes and oligonucleotides such as DNA. The first involves covalent interactions between the Lewis acidic metal and nucleic acid Lewis bases, such as nucleophilic guanine N7 residues. The second interaction occurs through intercalation of a small molecule between stacked base pairs of the double helix structure. Both such interactions cause disruption of base stacking in the double helix through interference with van der Waals or hydrogen-bonding interactions, and thus result in destabilization of double-helix conformation. The third interaction is electrostatic, occurring between a cationic complex and the anionic phosphate backbone.2,8,10 Neutralization of this negative charge reduces repulsive forces between adjacent phosphate groups, thus stabilizing the double helix.11,12 Of note is the fact that the former two interactions are often selective, while the electrostatic interactions may occur at any phosphate site along the phosphate backbone.13,14 These interactions can cause changes in the DNA structure, affecting cellular processes and potentially leading to cell death.4,8,10

10.3 Rhodium–DNA Interactions 10.3.1 Rhodium(I) Complexes Encouraged by the promising chemotherapeutic properties of cisplatin, a number of groups in the 1970s began to investigate the potential antineoplastic activity of other platinum group transition metal complexes, including rhodium. Preliminary work investigated the effects of cis-, square planar organometallic Rh(I) complexes of type [Rh(COD)L2]+ (L = 2,2′-bipyridine; 1,10-phenanthroline; piperidine) on growth of a number of tumours.15 In all cases, however, the Rh(I) complexes were less active than cisplatin. Similar observations were made on a series of related

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complexes.16 The reduced activity of such species was attributed to the instability of the Rh(I) centre towards oxidation to the kinetically inert octahedral Rh(III) complex in aqueous solution and therefore in vivo. In addition, the greatly enhanced lability of the leaving groups on Rh(I) versus Pt(II) complexes suggested that the Rh(I) species would be oxidized, and thus deactivated before reaching the target. No advanced studies on the interaction of Rh(I) species with oligonucleotides such as DNA were performed owing to these limitations.

10.3.2 Dirhodium(II) Complexes Dirhodium carboxylate complexes of general formula (Rh2(O2CR)4L2 (R = Me, Et, Pr; L = solvent) (Figure 10.1b) have been found to inhibit DNA replication and protein synthesis with a similar potency to cisplatin.17 In the examples given above, antitumour activity was found to increase with increasing lipophilicity of the R group, presumably through increased cell membrane permeability of the complex. Related complexes bearing various bidentate bridging species such as nitrogencontaining ligands18–20 (Figure 10.1c), phosphine ligands,21 (Figure 10.1d) and formamidinates22 (Figure 10.1e) have been developed, and exhibit antitumour activity comparable to, or higher than, cisplatin.17 Structure–activity relationships are, however somewhat more complex than the two-pole complimentary principle suggested by Yang et al. for organometallic antitumour agents.23 The similarities in antitumour properties between cisplatin derivatives and dirhodium complexes may be unexpected, owing to the obvious structural differences between the two species. Indeed the dirhodium complexes are capable of binding strongly to adenine residues, unlike cisplatin, which has a preference for guanine.17 Model experiments employed nucleo(s)tides to demonstrate that the interaction between rhodium and adenine residues is stabilized by favourable hydrogen bonds between the purine exocyclic NH2(6) group and a carboxylate oxygen atom of the dirhodium complex. In contrast, axial coordination of guanine residues is inhibited by electrostatic repulsions between the ketone O6 and oxygen atoms of the carboxylate ligands present within the complex.17,24 In subsequent studies, the unlikely substitution of two bridging ligands on complexes of type [Rh2(O2CR)4L2] by two guanine residues was directly observed by X-ray crystallographic analysis.25,26 The guanine residues were found to form equatorial bridging interactions, through the N7/O6 sites in a headto-head (H-H) (e.g. cis-[Rh2-(m-O2CCH3)2(9-EtGuaH)2(Me2CO)(H2O)](BF4)2 (Figure 10.1f)26 or head-to-tail (H-T) orientation (e.g. cis-[Rh2-(m-O2CCF3)2(9EtGuaH)2(Me2CO)2](CF3CO2)2 (Figure 10.1g).25 These findings led to further studies on interactions between dirhodium complexes and the dinucleotide d(GpG), where a similar chelate N7/O6 interaction with Rh2(OAc)4 was observed.27,28 Although the dirhodium complexes were found to bind through both axial and equatorial positions, the mechanism of binding is believed to occur through initial interaction through axial sites, followed by rearrangement to a more thermodynamically stable equatorial position.29,30 Indeed, recent studies demonstrated that placing nonlabile ligands in the axial position resulted in a decrease in the ability of the complexes to inhibit transcription in vitro, demonstrating the necessity for labile axial ligands, and

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Figure 10.1 Dinuclear Rh complexes

supporting the proposed rearrangement mechanism.31 The introduction of labile equatorial ligands may also facilitate ligand rearrangement to the equatorial position, and may therefore provide an explanation of the increased activity observed through introduction of electron-withdrawing groups, such as in Rh2(m-O2CF3)4, that exhibited activity comparable to that of cisplatin.32 In an extension to this work, dirhodium interactions with single-stranded oligonucleotides containing GG sites were investigated. This work further corroborated the correlation between lability of the ligands present and nucleic acid binding affinity, wherein the affinity of the dirhodium complexes were compared to cisplatin derivatives: cis-[Pt(NH3)2(H2O)2]2+ ≈ Rh2(m-O2CCF3)4 > cis-[Pt(NH3)2Cl2] >> cis-[Rh2(m-O2CCH3)2(NCCH3)6](BF4)2 > Rh2(mO2CCH3)4.17,33,34 More recent work has refuted the conclusions of earlier studies that suggested dirhodium complexes did not interact with double-stranded DNA. Indeed, interaction of a number of species demonstrated the presence of covalently linked DNA adducts. Significantly, these studies demonstrated the formation of stable DNA interstrand crosslinks35 Although specific binding has yet to be ascertained, the presence of a mixture of mono- and bifunctional adducts, including the more stable equatorial adducts was observed. Once again, the binding modes determined in model studies corroborated with data from the studies performed on doublestranded DNA, wherein an increased lability of the leaving groups corresponds to an increase in occurrence of interstrand crosslinks.36 As observed for binding of guanine residues to dirhodium complexes, bidentate equatorial binding of adenine residues on dinucleotide d(ApA) was observed, with the d(ApA) fragment

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spanning the Rh–Rh bond via N7/N6 bridges.33,37 Such observations again demonstrate the potential for dirhodium complexes to form interstrand crosslinks. The ability of dirhodium complexes to bind either adenine or guanine residues on plasmid pUC19 DNA was recently demonstrated, further disproving previous conclusions concerning the inability of such species to bind double-stranded DNA.38

10.3.3 Rhodium Complexes in Photodynamic Therapy Owing to the d-electron availability and interchangeable redox properties of transition metals, such complexes have found application in the area of photodynamic therapy (PDT).39 The DNA toxicity of oxygen-derived intermediates from PDT are well established, and light-induced formation of 1O2, O2− and •OH in the vicinity of DNA results in considerable degradation of DNA structure. The development of novel photoreagents based on bimetallic rhodium complexes has received considerable attention, owing to the propensity of such species to induce metal–metal charge transfer excited states to promote DNA cleavage. Dunbar and coworkers investigated the complex Rh2(m-O2CCH3)4, for potential photocleavage of DNA (Figure 10.2a).40 The d7-d7 Rh(II)-Rh(II) core was found to promote photoinduced cation formation in the presence of an electron acceptor. Subsequent DNA ‘nicking’ then occurred.41 Following these observations, a series of potential photoactivated complexes designed to interact with DNA both through coordination and intercalation were developed (Figure 10.2b).40,42,43 The complexes studied, with the exception of [Rh2(mO2CCH3)2(CH3CN)6](BF4)2,43 consisted of a chelating intercalating ligand on one of the two rhodium centres, with labile cis ligands on the second to facilitate covalent coordination to DNA nucleobases. Such species demonstrated DNA nicking upon photolysis, and cytotoxicity towards human skin cell lines.42 The dual binding capability of the related complex cis-[Rh2(dap)(m-O2CCH3)2(h1O2CCH3)(CH3OH)](O2CCH3) (dap = 1,12-diazaperylene) (Figure 10.2c),44 through both coordination and intercalation interactions with DNA, was elegantly demonstrated using 2D NMR studies.45 Similar studies were recently performed on a complex with two stable formamidinate and two labile trifluoroacetate bridging groups.37 cis-Rh2(DTolF)2(O2CCF3)2(H2O)2 (DTolF = N,N′-di-p-tolylformamidinate) (Figure 10.1e) has exhibited antitumour activity comparable to that of cisplatin against Yoshida ascites and T8 sarcomas, with a significant reduction in toxicity, showing virtually no toxicity in mice.22 Other heterobimetallic photodynamic species, bearing a Rh–Ru core have been developed,46 in which a low energy Ru–Rh metal–metal charge transfer excited state results in direct DNA nicking in the absence of electron acceptors. The extensive and elegant work performed by Dunbar and others has shed light on the potential mechanisms of activity of the structurally diverse dirhodium complexes. Despite the obvious differences in structure and reactivity between mononuclear cisplatin derivatives and the bimetallic rhodium complexes investigated, such research has demonstrated that the mode of activity may indeed be through formation of interstrand crosslinks between the rhodium metal centres and adjacent guanine or adenine residues on DNA. Through increased understanding

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Figure 10.2 Dinuclear rhodium complexes for photodynamic therapy

of the interactions between dirhodium species and DNA, novel complexes have been designed to bind DNA through both coordinative and intercalative interactions. Dimeric rhodium complexes have often demonstrated antitumour activity in the same order of magnitude as cisplatin, while potentially circumventing the toxicity issues associated with the successful drug. Furthermore, research into the potential of dirhodium complexes in photodynamic therapy may result in replacements for cisplatin derivatives in the foreseeable future.17–38, 40–46 10.3.4 Rhodium(III) Complexes The kinetic inertness of rhodium(III) complexes may be believed to preclude the ability of such species to modify or probe DNA and RNA function through covalent interactions. Indeed, divalent Pt, divalent (and trivalent) Ru species have received considerable attention in this field, directly owing to the lability and substitution chemistry of their ligands. Rhodium(III) complexes are low in the list of potential DNA interacting agents, owing to their inability to undergo substitution reactions on a physiologically relevant timescale.47 Further application is also limited by the inability of Rh(III), in contrast to Ru(III), to undergo activation through reduction to the divalent species. As a consequence, few cytotoxic Rh(III) complexes have been developed.8,10,48 A number of Rh(III) analogues of known antitumour active Ru(III) species have also been investigated. Like fac-[RuCl3(NH3)3], mer[RhCl3(NH3)3] is active, but insoluble.8 Rh(III) analogues of the active Ru(III) species, Na[trans-[RhCl4(Me2SO)(Im)] and (HIm)trans-[RhCl4(Im)2] were essentially inactive, exhibiting modest inhibition on the growth of the primary MCa mammary tumours.49 The limited activity in each case has been attributed to the kinetic stability of the Rh(III) relative to the Ru(III) systems towards ligand substitution.

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The concept of ‘photochemically’ activated Rh(III) cisplatin analogues has been investigated for complexes of the type cis-[RhCl2(polypyridyl)2]+.50,51 For example, cis-[RhCl2(phen)2]+ (phen 1,-10-phenanthroline) was found to undergo light-induced covalent binding to DNA.52 Although kinetically inert under thermal conditions, irradiation of the complexes results in d-electron promotion into ligandfield bands, thus increasing the rate of ligand exchange by orders of magnitude relative to thermal conditions. The reaction with N7 of guanine residues occurred owing to reductive quenching of the excited state by nucleobase–metal charge transfer, although with minimal biological effects.50 Such promising preliminary results led to the investigation of other complexes that demonstrated oxidative DNA strand scission.51 In particular, work by Barton and coworkers have led to many promising developments in this field. 10.3.5 Site-Specific Rhodium–DNA Interactions Barton and coworkers have produced a large body of work investigating the sitespecific interaction of Rh(III) intercalating complexes with DNA, with a view to introducing selective cleavage of the DNA strand in specific regions towards ‘sitedirected’ photodynamic therapy. Such work has also led to insights into the functions of nucleic acids in biological processes, and the complexes developed have been used both as photoactivated chemotherapeutic agents,51 and as probes for charge transport processes in DNA.53 The pairing of nucleobases in healthy cells occurs through combination of GC and AT residues. DNA base mismatches (i.e. CC, CA or CT combinations) arise owing to enzymatic errors or DNA damage during the process of genetic recombination or DNA replication.54,55 The inability of cells to repair mismatches is associated with cancerous transformations.56 Early work by Barton and coworkers involved the development of phenanthrenequinone diimine (phi) complexes of rhodium, that may intercalate into the major groove of DNA, and subsequently promote photoactivated strand scission.57–59 In an extension of this work, bulky heterocyclic aromatic imine ligands were introduced in complexes such as [Rh(bpy)2(chrysi)3+] (chrysi = 5,6chrysenequinonediimine, Figure 10.3).60 The bulky four-ring chrysene ligand, being too large to intercalate into standard DNA grooves, instead intercalated preferentially into the larger, perturbed site of the thermodynamically destabilized

Figure 10.3 [Rh(bpy)2(chrysi)3+] (chrysi = 5,6-chrysenequinonediimine)

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mismatch. Indeed, a direct correlation was observed between the extent to which a mismatch is destabilized (which depends on the bases involved) and the binding affinity of the complex for that mismatch.61 The complexes exhibited remarkably specific mismatch recognition, with photoactivated DNA cleavage observed at over 80% of mismatch sites, irrespective of the sequence of the mispaired bases involved.62 Significantly, cleavage was observed at a single-base mismatch in a 2725 base pair plasmid heteroduplex.61 Such high specificity has led to many applications of such complexes, including investigations into the correlation between deficiencies in mismatch repair and cancerous transformations, and towards the development of novel cancer treatments.56 Related complexes have also been employed as probes for mismatch occurrence;63,64 as targeting agents for conventional cisplatin chemotherapeutic protocols;65,66 and as site-specific delivery agents for metal-binding peptide conjugates.67 The complexes developed also inhibit cellular proliferation in mismatch-repairdeficient cells (such as those associated with cancerous growth) in preference to healthy, mismatch-proficient cells.68 Such observations have led to complexes designed for selective chemotherapeutic agents that target mismatches in cancerous, rather than healthy cells.68 Recent work has determined the mode of DNA binding involved in mismatch recognition. The crystal structure of [Rh(bpy)2(chrysi)]3+ bound to a CA mismatch in a DNA oligonucleotide revealed that, in contrast to earlier reports on the mode of intercalation, wherein DNA does not unwind to allow intercalation of the new ligand, the complex inserted into the double helix specifically at the mismatch site, ejecting both mismatched bases, to accommodate the new ligand.69 Such results again corroborate the ability of the bulky complex to specifically recognize mismatches over normal base pairing. Very recently the intercalation of a complex into a DNA cytosine–cytosine mismatch was identified using NMR solution studies.70

10.4 Tin–DNA Interactions 10.4.1 Introduction Tin complexes, in particular organotin species, where a direct C–Sn bond is present within the complex, have found application in many industries. The widespread use of organotin complexes in the food, agricultural, paints and polymer industries has resulted in their accumulative distribution in the environment and in biological systems. The biological toxicity of tin species is well known, and such species exhibit a particular toxicity towards the central nervous system. The ability of inorganic divalent tin to interact with DNA has been established.71–73 Such species were found to enter human white blood cells, and cause extensive damage to DNA through oxidative mechanisms. As a consequence, much research has focused on the biological effects of organotin complexes.2,8,23,74,75 Despite the associated toxicity issues, organotin compounds have found many pharmacological applications, as potential antiinflammatory, antimicrobial and antitumour agents. Understanding the interac-

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tions of organotin reagents with biological species such as DNA is therefore a principle area of research in preventing, and exploiting, the toxicity of tin complexes.

10.4.2 Organotin Complexes–DNA Interactions Organotin(IV) complexes of the type X2R2Sn and X2R2L2Sn (X = halide, pseudohalide; R = alkyl, aryl; L = nitrogen ligand, e.g. py, bpy) have demonstrated moderate activity against a range of tumours.8 The application of such species in the clinic, however, is limited by their toxicity, which has been found to increase in the order RSn < R2Sn < R3Sn. An increase in the lipophilicity of the R group has been correlated with an increase in toxicity, and alkyl complexes are often more toxic than their aryl analogues. The principle forms of toxicity are neurological, and may occur as a result of the affinity of tin for sulfur, i.e. through coordination to protein thiols.76 The correlation between lipophilicity and toxicity may be related to the cell membrane-permeability of lipophilic species. Maintaining a balance between the hydrophobic and hydrophilic character of the complexes involved, therefore, has been a focus of much research, with the intention of increasing antitumour activity, while reducing toxicity issues. For example, highly lipophilic stannocene-type complexes such as decaphenyl stannocene, [h5-C5(C6H5)5]2Sn, structurally analogous to the antitumour active titanocene dichloride,2 have exhibited antitumour activity against Ehrlich Ascites tumour with cure rates of 68–90%, remarkably with relatively low toxicity.77 The fact that such sterically crowded Sn(II) centres exhibit antitumour activity points towards a different mode of activity than covalent coordination of these complexes to nucleobases on DNA. Although not discussed by the authors, potential intercalation of the planar aromatic substituents on the stannocenes into DNA (see above) may be a potential mechanism of activity, which may also explain why the pentasubstituted cyclopentadienyl ligands exhibit some antitumour activity, albeit less pronounced than the corresponding stannocene. While the low water solubility of the majority of organotin compounds is a significant problem, the introduction of hydrophilic leaving groups8,74,75 or, for example, their inclusion into b-cyclodextrin75 may circumvent such limitations. In contrast to cisplatin, the mechanism of activity of organotin compounds has not been well established. Owing to the structural similarity of such complexes to cisplatin, DNA has been frequently proposed as a potential therapeutic target and, in order to gain further understanding on the antitumour activity of such species, the interactions of organotin complexes with DNA have been investigated by a number of groups. Organotin(IV) complexes have been shown to coordinate to bis(adenine) ligands to form tetrahedral adducts. Reactions of guanine, thymine, cytosine and uracil under similar conditions, however, afforded no coordination products.78 Indeed, the interaction of diorganotin(IV) compounds such as R2SnCl2(phen) (R = Et, n-Bu, Ph) with nucleotide monophosphates in neutral aqueous solution afford ill-defined, polymeric species in which the Sn(IV) centre coordinates only to the phosphate groups, with no observed coordination to the nucleobases present.23,75,79 Interaction of Bu2SnCl2 or Bu3SnCl with 5′-AMP and 5′-GMP exhibited multiple

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Figure 10.4 Organotin–DNA adduct formation

binding modes to the Sn(IV) centre through 31P NMR analysis, wherein the Sn(IV) centre could be bound in a chelate or nonchelate manner to the phosphate oxygen.80 In 119Sn Mossbauer studies, Barbieri observed the interaction of solvated Sn(IV) species (R2SnCl2(EtOH)2 and R3SnCl(EtOH), R= Me, Et, n-Bu, n-Oct or Ph), in which possible DNA adducts, R2Sn(DNA)2 and R3Sn(DNA) were observed as gel phases (DNA condensation).81–83 Adduct formation was proposed to occur by neutralization of the anionic phosphate groups by the cationic Sn centre, with proposed octahedral and trigonal bipyramidal structures (Figure 10.4). Interestingly, while the active species for cisplatin antitumour activity is known to be the hydrolysis product, ([(NH3)2Pt(OH2)2]2+) analogous Sn species, [Me2Sn(OH)(H2O)n]+, Me2Sn(OH)2 and Me3Sn(OH)(H2O)2 did not interact with native DNA.84 Such observations led the authors to propose structure–activity relationships, wherein the ability of organotin(IV) complexes to bind DNA depends on the lipophilicity of the complex. Therefore, as lipophilicity increases for RnSn(IV) in the series: R = Me < Et < Ph < n-Bu < n-Oct, so the tendency of the complex to form DNA-condensates increases.82 When formation of DNA condensates does occur, the interaction was proposed to occur through electrostatic interaction between ethanol solvated species and oxygen atoms of the phosphodiester groups present. Later theoretical and model studies on organotin(IV)–DNA interactions further supported the conclusions that binding of such species occurs through electrostatic interaction with phosphate O atoms.85,86 Binding of organotin(IV) species to O donor ligands in nucleotides has exhibited pH dependence in a number of studies. For example, coordination of hydrated [Me2Sn(IV)]2+ with a series of nucleotides under acidic conditions (pH < 4) resulted in binding through phosphate oxygen atoms. At intermediate pH (4–9.5), no interaction occurred, while under basic conditions, (pH > 9.5) the hydroxyl groups of the sugar functionality present on the nucleotide were also capable of coordination to the Sn centre, in particular if a second deprotonated hydroxyl group was adjacent to the first, to form a chelate complex.8 Similar observations were made in elegant studies on the interaction of Et2SnCl2(phen) with 5′-dGMP, using [trans-en2Os(nH2)](CF3SO3)2 as 1H-NMR probe.8,75 In all cases, the pH dependence of the Sn–DNA interaction was observed. The chemical and structural differences between organotin complexes and cisplatin would suggest that very different modes of antitumour activity are exhibited by each. The principle sites of interaction for organotin species would appear

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categorically to be oxygen atoms of phosphate and, depending on pH, hydroxyl groups on the ribose or deoxyribose moieties present. Further studies on the interaction of organotin(IV) complexes with calf thymus DNA revealed that no disruption of DNA double helix structure occurs. Neither does any DNA degradation occur through oxidative stress mechanisms.87 The question of how organotin complexes may inhibit tumour growth remains, as yet, unanswered.

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20. Pruchnik, F.; Dus, D.; Properties of rhodium(II) complexes having cytostatic activity. J. Inorg. Biochem., 1996, 61(1), 55–61. 21. Pruchnik, F.P.; Starosta, R.; Ciunik, Z.; Opolski, A.; Wietrzyk, J.; Wojdat, E. et al.; Tetraacetatodirhodium(II) complexes with tris(methoxyphenyl) phosphines, their reactivity, structure, and antitumor activity; Can. J. Chem., 2001,79(5/6), 868–877. 22. Fimiani, V.; Ainis, T.; Cavallaro, A.; Piraino, P.; Antitumor effect of the new rhodium(II) complex: Rh2(Form)2(O2CCF3)2(H2O)2 (Form = N,N′-di-p-tolyl formamidinate); J. Chemother., 1990, 2(5), 319–326. 23. Yang, P.; Guo, M.; Interactions of organometallic anticancer agents with nucleotides and DNA; Coord. Chem. Rev., 1999, 185–186, 189–211. 24. Aoki, K.; Salam, M.A.; Interligand interactions affecting specific metal bonding to nucleic acid bases. A case of [Rh2(OAc)4], [Rh2(HNOCCF3)4], and [Rh2(OAc)2(HNOCCF3)2] toward purine nucleobases and nucleosides; Inorg. Chim. Acta, 2002, 339, 427– 437. 25. Dunbar, K.R.; Matonic, J.H.; Saharan, V.P.; Crawford, C.A.; Christou, G.; Structural evidence for a new metal-binding mode for guanine bases: Implications for the binding of dinuclear antitumor agents to DNA; J. Am. Chem. Soc., 1994, 116(5), 2201–2202. 26. Crawford, C.A.; Day, E.F.; Saharan, V.P.; Folting, K.; Huffman, J.C.; Dunbar, K.R. et al.; N7,O6 bridging 9-ethylguanine (9-EtGH) groups in dinuclear metal-metal bonded complexes with bond orders of one, two or four; J. Chem. Soc. Chem. Comm., 1996, (10), 1113–1114. 27. Chifotides, H.T.; Koshlap, K.M.; Perez, L.M.; Dunbar, K.R.; Novel binding interactions of the DNA fragment d(pGpG) cross-linked by the antitumor active compound tetrakis(mcarboxylato)dirhodium(II,II); J. Am. Chem. Soc., 2003, 125(35), 10714–1024. 28. Chifotides, H.T.; Koshlap, K.M.; Perez, L.M.; Dunbar, K.R.; Unprecedented head-to-head conformers of d(GpG) bound to the antitumor active compound tetrakis(m-carboxylato )dirhodium(II,II); J. Am. Chem. Soc., 2003, 125(35), 10703–10713. 29. Perlepes, S.P.; Huffman, J.C.; Matonic, J.H.; Dunbar, K.R.; Christou, G.; Binding of 2,2′bipyridine to the dirhodium(II) tetraacetate core: unusual structural features and biological relevance of the product Rh2(OAc)4(bpy); J. Am. Chem. Soc., 1991, 113(7), 2770–2771. 30. Crawford, C.A.; Matonic, J.H.; Streib, W.E.; Huffman, J.C.; Dunbar, K.R.; Christou, G.; Reaction of 2,2′-bipyridine (bpy) with dirhodium carboxylates: mono-bpy products with variable chelate binding modes and insights into the reaction mechanism; Inorg. Chem., 1993, 32(14), 3125–3133. 31. Aguirre, J.D.; Lutterman, D.A.; Angeles-Boza, A.M.; Dunbar, K.R.; Turro, C.; Effect of axial coordination on the electronic structure and biological activity of dirhodium(II,II) complexes; Inorg. Chem., 2007, 46(18), 7494–7502. 32. Reibscheid, E.M.; Zyngier, S.; Maria, D.A.; Mistrone, R.J.; Sinisterra, R.D.; Couto, L.G. et al.; Antitumor effects of rhodium (II) complexes on mice bearing Ehrlich tumors; Braz. J. Med. Biol. Res., 1994, 27(1), 91–94. 33. Chifotides, H.T.; Koomen, J.M.; Kang, M.; Tichy, S.E.; Dunbar, K.R.; Russell, D.H.; Binding of DNA purine sites to dirhodium compounds probed by mass spectrometry; Inorg. Chem., 2004, 43(20), 6177–6187. 34. Asara, J.M.; Hess, J.S.; Lozada, E.; Dunbar, K.R.; Allison, J.; Evidence for binding of dirhodium bis-acetate units to adjacent GG and AA sites on single-stranded DNA; J. Am. Chem. Soc., 2000, 122(1), 8–13. 35. Dunham Shari, U.; Chifotides Helen, T.; Mikulski, S.; Burr Amity, E.; Dunbar, Kim R.; Covalent binding and interstrand cross-linking of duplex DNA by dirhodium(II,II) carboxylate compounds; Biochemistry, 2005, 44(3), 996–1003. 36. Chifotides, H.T.; Fu, P.K.L.; Dunbar, K.R.; Turro, C.; Effect of equatorial ligands of dirhodium(II,II) complexes on the efficiency and mechanism of transcription inhibition in vitro; Inorg. Chem.; 2004, 43(3), 1175–1183. 37. Chifotides, H.T.; Dunbar, K.R.; Unprecedented head-to-head right-handed cross-links between the antitumor bis(m-N,N′-di-p-tolylformamidinate) dirhodium(II,II) core and

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the dinucleotide d(ApA) with the adenine bases in the rare imino form; J. Am. Chem. Soc.; 2007, 129(41), 12480–12490. Rahman, M.M.; Yasuda, H.; Katsura, S.; Mizuno, A.; Inhibition of endonuclease cleavage and DNA replication of E. coli plasmid by the antitumor rhodium(II) complex; Arch. Biochem. Biophys., 2007, 464(1), 28–35. Detty, M.R.; Gibson, S.L.; Wagner, S.J.; Current clinical and preclinical photosensitizers for use in photodynamic therapy; J. Med. Chem.; 2004, 47(16), 3897–3915. Bradley, P.M.; Angeles-Boza, A.M.; Dunbar, K.R.; Turro, C.; Direct DNA photocleavage by a new intercalating dirhodium(II/II) complex: comparison to Rh2(m-O2CCH3)4; Inorg. Chem., 2004, 43(8), 2450–2452. Fu, P.K.L.; Bradley, P.M.;Turro, C.; DNA cleavage by photogenerated Rh2(O2CCH3)4(H2O)2+; Inorg. Chem.; 2001, 40(11), 2476–2477. Angeles-Boza, A.M.; Bradley, P.M.; Fu, P.K.L.; Wicke, S.E.; Bacsa, J.; Dunbar, K.R. et al.; DNA binding and photocleavage in vitro by new dirhodium(II) dppz complexes: correlation to cytotoxicity and photocytotoxicity; Inorg. Chem.; 2004, 43(26), 8510–8519. Lutterman, D.A.; Fu, P.K.L.; Turro, C.; cis-[Rh2(m-O2CCH3)2(CH3CN)6]2+ as a photoactivated cisplatin analog; J. Am. Chem. Soc., 2006, 128(3), 738–739. Chouai, A.; Wicke, S.E.; Turro, C.; Bacsa, J.; Dunbar, K.R.; Wang, D. et al.; Ruthenium(II) complexes of 1,12-diazaperylene and their interactions with DNA; Inorg. Chem., 2005, 44(17), 5996–6003. Kang, M.; Chouai, A.; Chifotides, H.T.; Dunbar, K.R.; 2D NMR spectroscopic evidence for unprecedented interactions of cis-[Rh2(dap)(m-O2CCH3)2(H1-O2CCH3)(CH3OH)] (O2CCH3) with a DNA oligonucleotide: combination of intercalative and coordinative binding; Angew. Chem., Int. Ed., 2006, 45(37), 6148–6151. Swavey, S.; Brewer, K.J.; Visible light induced photocleavage of DNA by a mixed-metal supramolecular complex: [{(bpy)2Ru(dpp)}2RhCl2]5+; Inorg. Chem., 2002, 41(24), 6196–6198. Barry, C.G.; Turney, E.C.; Day, C.S.; Saluta, G.; Kucera, G.L.; Bierbach, U.; Thermally inert metal ammines as light-inducible DNA-targeted agents. synthesis, photochemistry, and photobiology of a prototypical rhodium(III)-intercalator conjugate; Inorg. Chem., 2002, 41(26), 7159–7169. Pieper, T.; Borsky, K.; Keppler, B.K.; Non-platinum antitumor compounds. Topics Biol. Inorg. Chem., 1999, 1(Metallopharmaceuticals I),171–199. Mestroni, G.; Alessio, E.; Sessanta o Santi, A.; Geremia, S.; Bergamo, A.; Sava, G. et al.; Rhodium(III) analogs of antitumor-active ruthenium(III) compounds: the crystal structure of [ImH][trans-RhCl4(Im)2] (Im = imidazole); Inorg. Chim. Acta, 1998, 273(1,2), 62–71. Billadeau, M.A.; Morrison, H.; Photolytic covalent binding of metal complexes to DNA. In: Metal ions in Biological Systems, Vol. 33, Sigel, A. and Sigel, H., Eds., Marcel Dekker, New York, 1996, 269–296. Erkkila, K.E.; Odom, D.T.; Barton, J.K.; Recognition and reaction of metallointercalators with DNA; Chem. Rev., 1999, 99(9), 2777–2795. Mahnken, R.E.; Billadeau, M.A.; Nikonowicz, E.P.; Morrison, H.; Development of photo cis-platinum reagents. Reaction of cis-dichlorobis(1,10-phenanthroline)rhodium(III) with calf thymus DNA, nucleotides and nucleosides; J. Am. Chem. Soc., 1992, 114(24), 9253–9265. O’Neill, M.A.; Barton, J.K.; DNA-mediated charge transport chemistry and biology; Topics Curr. Chem., 2004, 236 (Long-Range Charge Transfer in DNA I), 67–115. Modrich, P.; Mechanisms and biological effects of mismatch repair; Ann. Rev. Genet., 1991, 25, 229–253. Kolodner, R.; Biochemistry and genetics of eukaryotic mismatch repair; Genes Devel., 1996, 10(12), 1433–1442. Kunkel, T.A.; Erie, D.A.; DNA mismatch repair; Ann. Rev. Biochem., 2005, 74, 681–710. David, S.S.; Barton, J.K.; NMR evidence for specific intercalation of D-Rh(phen)2phi3+ in [d(CTCGAC)2]; J. Am. Chem. Soc., 1993, 115(7), 2984–2985.

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58. Hudson, B.P.; Dupureur, C.M.; Barton, J.K.; 1H NMR structural evidence for the sequence-specific design of an intercalator: ∆-a-[Rh[(R,R)-Me2trien]phi]3+ bound to d(GAGTGCACTC)2; J. Am. Chem. Soc., 1995, 117(36), 9379–9380. 59. Sitlani, A.; Long, E.C.; Pyle, A.M.; Barton, J.K.; DNA photocleavage by phenanthrenequinone diimine complexes of rhodium(III): shape-selective recognition and reaction; J. Am. Chem. Soc., 1992, 114(7), 2303–2312. 60. Jackson, B.A.; Barton, J.K.; Recognition of DNA base mismatches by a rhodium intercalator; J. Am. Chem. Soc., 1997, 119(52), 12986–12987. 61. Jackson, B.A., Barton, J.K.; Recognition of base mismatches in DNA by 5,6-chrysenequinone diimine complexes of rhodium(III): A proposed mechanism for preferential binding in destabilized regions of the double helix; Biochemistry; 2000, 39(20), 6176–6182. 62. Jackson, B.A.; Alekseyev, V.Y.; Barton, J.K.; A versatile mismatch recognition agent: specific cleavage of a plasmid DNA at a single base mispair; Biochemistry, 1999 38(15), 4655–4662. 63. Rueba, E.; Hart, J.R.; Barton, J.K.; [Ru(bpy)2(L)]Cl2: Luminescent metal complexes that bind DNA base mismatches; Inorg. Chem., 2004, 43(15), 4570–4578. 64. Zeglis, B.M.; Barton, J.K.; A mismatch-selective bifunctional rhodium-oregon green conjugate: a fluorescent probe for mismatched DNA; J. Am. Chem. Soc., 2006, 128(17), 5654–5655. 65. Schatzschneider, U.; Barton, J.K.; Bifunctional rhodium intercalator conjugates as mismatch-directing DNA alkylating agents; J. Am. Chem. Soc., 2004, 126(28), 8630–8631. 66. Petitjean, A.; Barton, J.K.; Tuning the DNA reactivity of cis-platinum: Conjugation to a mismatch-specific metallointercalator; J. Am. Chem. Soc., 2004, 126(45), 14728–14729. 67. Copeland, K.D.; Fitzsimons, M.P.; Houser, R.P.; Barton, J.K.; DNA hydrolysis and oxidative cleavage by metal-binding peptides tethered to rhodium intercalators; Biochemistry, 2002, 41(1), 343–356. 68. Hart, J.R.; Glebov, O.; Ernst, R.J.; Kirsch, I.R.; Barton, J.; DNA mismatch-specific targeting and hypersensitivity of mismatch-repair-deficient cells to bulky rhodium(III) intercalators; Proc. Natl. Acad. Sci. USA, 2006, 103(42), 15359–15363. 69. Pierre, V.C.; Kaiser, J.T.; Barton, J.K.; Insights into finding a mismatch through the structure of a mispaired DNA bound by a rhodium intercalator; Proc. Natl. Acad. Sci. USA, 2007, 104(2), 429–434. 70. Cordier, C.; Pierre, V.C.; Barton, J.K.; Insertion of a bulky rhodium complex into a DNA cytosine-cytosine mismatch: An NMR solution study; J. Am. Chem. Soc., 2007, 129(40), 12287–12295. 71. McLean, J.R.N.; Birnboim, H.C.; Pontefact, R.; Kaplan, J.G.; The effect of tin chloride on the structure and function of DNA in human white blood cells; Chem. Biol. Inter., 1983, 46(2), 189–200. 72. McLean, J.R.N.; Blakey, D.H.; Douglas, G.R.; Kaplan, J.G.; The effect of stannous and stannic (tin) chloride on DNA in Chinese hamster ovary cells; Mut. Res. Lett., 1983, 119(2), 195–201. 73. Dantas, F.J.S., Moraes, M.O.; Carvalho, E.F.; Valsa, J.O.; Bernardo-Filho, M.; Caldeira-DeAraujo, A.; Lethality induced by stannous chloride on Escherichia coli AB1157: participation of reactive oxygen species; Food Chem. Toxicol., 1996, 34(10), 959–962. 74. De Vos, D.; Willem, R.; Gielen, M.; Van Wingerden, K.E.; Nooter, K.; The development of novel organotin antitumor drugs: structure and activity; Metal-Based Drugs. 1998, 5(4), 179–188. 75. Pellerito, L.; Nagy, L.; Organotin(IV)n+ complexes formed with biologically active ligands: equilibrium and structural studies, and some biological aspects; Coord. Chem. Rev., 2002, 224(1–2), 111–150. 76. Penninks, A.H.; Seinen, W.; Mechanisms of dialkyltin-induced immunopathology; Vet. Quart., 1984, 6(4), 209–215. 77. Kopf-Maier, P.; Janiak, C.; Schumann, H.; Antitumor properties of organometallic metallocene complexes of tin and germanium; J. Cancer Res. Clin. Oncol., 1988, 114(5), 502–506.

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78. Cardin, C.J.; Roy, A.; Anticancer activity of organotin compounds. 2. Interaction of diorganotin dihalides with nucleic acid bases and nucleosides; the synthesis of adenine, adenosine and 9-methyladenine adducts; Inorg. Chim. Acta, 1985, 107(1), 57–61. 79. Barbieri, R.; Silvestri, A.; Piro, V.; Tin-119 Moessbauer titration of dimethyl- and trimethyltin(IV) hydroxides with model ligands mimicking nucleic acid phosphate sites, and with deoxyribonucleic acid; J. Chem. Soc., Dalton Trans.: 1990, (12), 3605–3609. 80. Atkinson, A.; Rodriguez, M.D.; Shewmaker, T.E.; Walmsley, J.A.; Synthesis and characterization of compounds of di- and tributyltin chloride with adenine and guanine mononucleotides; Inorg. Chim. Acta, 1999, 285(1), 60–69. 81. Barbieri, R.; Silvestri, A.; Giuliani, A.M.; Piro, V.; Di Simone, F.; Madonia, G.; Organotin compounds and deoxyribonucleic acid; J. Chem. Soc., Dalton Trans.: 1992, (4), 585–590. 82. Piro, V.; Di Simone, F.; Madonia, G.; Silvestri, A.; Giuliani, A.M.; Ruisi, G. et al.; The interaction of organotins with native DNA; Appl. Organomet. Chem., 1992, 6(6), 537–542. 83. Barbieri, R.; Ruisi, G.; Silvestri, A.; Giuliani, A.M.; Barbieri, A.; Spina, G. et al.; Dynamics of tin nuclei in alkyltin(IV)-deoxyribonucleic acid condensates by variable-temperature tin-119 Moessbauer spectroscopy; J. Chem. Soc., Dalton Trans.: 1995, (3), 467–475. 84. Barbieri, R.; Silvestri, A.; The hydrolysis of Me2SnIV and Me3SnIV moieties monitored through tin-119 Moessbauer spectroscopy; Inorg. Chim. Acta, 1991, 188(1), 95–98. 85. Barone, G.; Ramusino, M.C.; Barbieri, R.; La Manna, G.; Semiempirical calculations on the interaction between dimethyltin(IV) and DNA model system; Theochem., 1999, 469, 143–149. 86. Barbieri, R.; Huber, F.; Silvestri, A.; Ruisi, G.; Rossi, M.; Barone, G. et al.; The interaction of S,N-coordinated dimethyltin(IV) derivatives with deoxyribonucleic acid: structure and dynamics by 119Sn Mossbauer spectroscopy; Appl. Organomet. Chem., 1999, 13(8), 595–603. 87. Barone, G.; Barbieri, R.; La Manna, G.; Koch, M.H.J.; The interaction of deoxyribonucleic acid with methyltin(IV) moieties in solution studied by small-angle X-ray scattering, circular dichroism and UV spectroscopy; Appl. Organomet. Chem. 2000, 14(4), 189–196.

Part C DNA Recognition: Nucleases and Sensors

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

11 Groove-Binding Ruthenium(II) Complexes as Probes of DNA Recognition Jayden A. Smith, J. Grant Collins and F. Richard Keene

11.1 Introduction The application of polypyridyl transition metal complexes to the structural elucidation of biological molecules had its origins in the observations of Dwyer et al. in the 1950s.1 The different biological activity of tris(phenanthroline) complexes of Fe(II), Ni(II), Ru(II), Os(II), Co(II) and Zn(II) in laboratory mice was reported, with selectivity of the D (right-handed) in preference to the L (left-handed) enantiomer noted in the case of inert species. They also noted a curariform activity and significant antibacterial characteristics, and a significant enhancement of activity on methyl substitution on the ligands in the cases of iron and ruthenium species. As such complexes were chemically inert, Dwyer et al. reasoned that any effect they had must be based primarily on their physical interactions with biological systems. Ruthenium has often been the metal of choice when studying the interactions of polypyridyl transition metal complexes with nucleic acids, owing to the wellestablished chemistry and rich photophysical properties of this genre.2 Polypyridylruthenium complexes are typically chemically inert, meaning their interactions with nucleic acids are usually noncovalent and hence reversible, and being octahedral tris(bidentate) species they possess an inherent chirality which can be exploited in their interactions with chiral polynucleotides. Furthermore, these complexes exhibit strong metal-to-ligand charge transfer (MLCT)-induced absorbances in the visible Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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Groove-Binding Ruthenium(II) Complexes

region of the spectrum and notable luminescent emissions – perturbations of these spectral characteristics upon nucleic-acid binding provide a convenient means to appraise the extent or nature of the interaction.

11.2 Mononuclear Complexes [Ru(phen)3]2+ (phen = 1,10-phenanthroline; see Figure 11.1d) has received particular attention as an inert metalloprobe of biological molecules. Its DNA binding ability has been most extensively investigated by Barton and coworkers, who in the early 1980s, elaborated upon the work of Dwyer et al. by investigating the in vitro enantioselectivity of the complex.3 On the basis of hypochromicity measurements it was proposed that the complex bound via intercalation of a single phenanthroline ligand. Preferential association between the D form of the complex and right-handed B-DNA was confirmed. The observed enantioselectivity is believed to be due to the unfavourable steric interactions between the nonintercalated ligands of the L enantiomer with the sugar-phosphate backbone of the DNA. Alternatively, the nonintercalating phenanthroline ligands of the D isomer fit easily within the complementary right-handed helical groove. No enantioselectivity was observed in experiments conducted with left-handed Z-DNA, presumably due to the altered groove dimensions. The intercalative model proposed by Barton et al. was supported by experiments utilizing the analogous complex with 4,7-diphenyl-1,10-phenanthroline (DIP; Figure 11.1o) as the ligand. As expected, the phenyl groups heightened the observed enantioselectivity, purportedly by exaggerating the steric interactions between the L enantiomer and the phosphate backbone, as well as enhancing the extent of intercalation.4 Again, both enantiomers were found to bind to Z-DNA with equal affinity. By contrast, [Ru(bpy)3]2+ (bpy = 2,2′-bipyridine; Figure 11.1a), lacking sufficiently elongated polyaromatic ligands to participate in intercalation, associated with DNA only weakly in what was believed to be primarily an electrostatic interaction.5 However, the intercalative binding proposal has been the subject of considerable controversy. Viscosity measurements undertaken by Chaires et al.6 and scanning force microscopy investigations by Williams, Bottomley et al.7 both suggested a nonintercalative mode of binding. Subsequent NMR experiments of Collins et al.8 gave some indication that [Ru(phen)3]2+ may indeed be partially intercalating, a result confirmed by more recent linear dichroism studies by Nordén and Lincoln:9 they claimed that both enantiomers of the complex may bind with one phenanthroline ligand partially (semi- or quasi-) intercalated via the minor groove, and their proposed binding geometry is in reasonable agreement with the available NMR data. This partial intercalation model is commonly seen amongst complexes which have polyaromatic ligands that either have insufficient span to insert fully between stacked base pairs, or in cases where the intercalating ligand deviates from planarity, inhibiting complete insertion.10 Indeed, a circular and linear dichroism study by Nordén and coworkers eventually dismissed an intercalative binding mode for [Ru(DIP)3]2+.11

Mononuclear Complexes

H3C

321

CH3

N

N

(a) bpy

N

N

(d) phen

(c) tBu2bpy

(b) Me2bpy

H3C

CH3

N

N

N

N

CH3

H3C

H3C

CH3

CH3

H3C N

N

H3C

N

CH3

N

N

N

N

(g) 4,7-Me2phen

(e) 2,9-Me2phen

N

(h) TMP

(f) 5,6-Me2phen

N

N

N

N

N

N

N

N

N

N

N

N

N

N

(l) dppx

(j) dppz

CH3

H3C

N H

N

(k) dpqC

(i) dpq

H2N

N

N H

N

NH

(o) DIP

NH2

(m) (R,R)-Me2trien

N

N N

N

(n) IP

HN

H3C N

(q) picchxnMe2

HN (p) phi

N

N

N H3C

N

(r) 2-appt

N H2N

Figure 11.1 Bidentate and tetradentate ligands

N N

N H

322

Groove-Binding Ruthenium(II) Complexes

(s) bbtb N

S N

S N

N

H N

N

N

Y

(t) PIP

N

N

N

NH

N

N

NH

(v) chrysi

(u) tactp

Figure 11.1 Continued

The controversy surrounding the exact mode of binding adopted by [Ru(phen)3]2+, heightened by the relatively low binding affinity of the complex (∼6 × 103 M−1, dependent upon DNA sequence),3,12 was exacerbated by debate over whether the complex was binding in the major or the minor groove. Two-dimensional NMR experiments have revealed that both enantiomers of [Ru(phen)3]2+ bind in the minor groove of DNA.13 The numerous conflicting results regarding the site and orientation of binding have been attributed to variations in technique, DNA sequence and even metal complex/DNA/salt concentrations.14,15 A minor-groove binding mode was consistent with the trend observed amongst the bulk of other polypyridylruthenium(II) complexes studied.16–18 The binding nature of the related complex [Ru(phen)2(dppz)]2+ (dppz = dipyrido[3,2-a:2′,3′-c] phenazine; Figure 11.1j) has also invoked vigorous debate.19 Complexes based on the dppz ligand function as ‘molecular light switches’: in aqueous solution [Ru(bpy)2(dppz)]2+ and [Ru(phen)2(dppz)]2+ are effectively quenched by the proton-donating solvent and show no luminescence, but when intercalated to DNA the hydrophobic environment allows them to luminesce strongly.20,21 Additionally, the facile electron transfer capabilities of intercalated ruthenium dppz complexes has seen their frequent use in studies of long-range DNA-mediated electron transfer.20,22–26 Unlike phen, the intercalative ability of the dppz ligand was unambiguous and undisputed, having been proven using a variety of techniques and methodologies.16,21,27,28 [Ru(phen)2(dppz)]2+ has been found to bind to DNA with an association constant in excess of 106 M−1:27,28 the extended planar aromatic ring system of the dppz ligand lends itself to intercalative stacking between base pairs much more readily than its less-extended phen counterpart.29–31 Nevertheless, there still exists a significant electrostatic contribution to the binding of dppz complexes.27

Mononuclear Complexes

323

Despite universal support for the intercalative binding mode of dppz-based complexes, the groove via which the complexes intercalated became a point of contention, but present understanding is that it binds from the minor groove. NMR studies have suggested that both D-[Ru(phen)2(dpq)]2+ (dpq = dipyrido[3,2-d:2′,3′-f] quinoxaline; Figure 11.1i) and D-[Ru(phen)2(dpqC)]2+ (dpqC = dipyrido[3,2-a:2′,3′-c](6,7,8,9-tetrahydro)phenazine; Figure 11.1k) bind via the minor groove.8,32 The dpq and dpqC complexes do not bind to DNA as strongly as do their dppz or dppx analogues (dppx = 7,8-dimethyldipyridophenazine; Figure 11.1l), presumably as they cannot intercalate as effectively as dppz or dppx, which possess larger aromatic areas that may potentially stack between DNA bases. Additional NMR experiments conducted by Collins et al. on [Ru(Me2phen)2(dpq)]2+ and [Ru(Me2phen)2(dppz)]2+ (2,9-Me2phen = 2,9-dimethyl-1,10-phenanthroline; Figure 11.1e) also suggested intercalation via the minor groove (methylated phenanthroline ligands were used to simplify the NMR spectra and to provide strong NOE signals in NOESY spectra).17,18 Recently, Nordén and coworkers conceded the possibility of intercalation from either groove when the other is blocked; this concession was based in part on spectroscopic investigations into the properties of DNA-bound [Ru(phen)2(dppz)]2+ in the presence and absence of the minor-groove binder 4′,6diamidino-2-phenylindole (DAPI).33,34 Negligible enantioselectivity is observed in the DNA-binding of [Ru(phen)2(dppz)]2+ 17,28 and its 2,2′-bipyridine analogue [Ru(bpy)2(dppz)]2+,34 although in each case differential luminescence and binding rates were observed between the D and L isomers of each complex, suggesting different binding geometries for each enantiomer. The bulk of studies conducted into polypyridyl complex–nucleic acid interactions have concerned themselves with intercalating mononuclear complexes, typically of the form [Ru(ancillary)2(intercalator)]2+, where there is a single dedicated intercalating ligand and a pair of ancillary ligands that occupy the groove (to a variable extent). Due to the three-dimensional nature of the association between these octahedral complexes and their chiral polynucleotide targets, the nature of all the ligands − ancillary or intercalating − influences the binding affinity of the complex although a larger intercalative surface area generally correlates with a greater binding affinity.35 Molecular modelling suggests that increasing the length of the intercalating ligand favours binding via the minor groove, whereas increases to the width of this ligand promote binding from the major groove.36 These findings are corroborated by experimental results obtained with Ru(II) and Rh(III) complexes featuring the relatively wide intercalating ligand phi (phenanthrenequinone diimine; Figure 11.1p). NMR experiments suggest that D-cis-a-[Ru(RR-picchxnMe2)(dpq)]2+ (picchxnMe2 = N,N′-dimethyl-N,N′-di(2-picolyl)-1,2-diaminocyclohexane; Figure 11.1q) intercalates via the minor groove of DNA, whereas the analogous complex with a phi intercalator, D-cis-a-[Ru(RR-picchxnMe2)(phi)]2+, binds via the major groove.37 The phi ligand has seen particularly extensive use in octahedral rhodium complexes, and such species almost invariably intercalate via the major groove.38 Rhodium complexes containing phi, or derivatives thereof, have demonstrated an impressive degree of selectivity for features such as base sequences,38,39 sequence-dependent

324

Groove-Binding Ruthenium(II) Complexes

twisting,40,41 base mismatches42,43 and base triples.44 The usual enantioselectivity favouring the D form is observed in DNA-binding experiments with these complexes.45 The role of ancillary ligands is therefore important, as the steric bulk of the nonintercalating ligands directly influences how deeply the complex may intercalate. In addition, hydrophobic interactions, van der Waals contacts, and hydrogen bonding brought about by the ancillary ligands are believed to be the major driving forces behind many DNA-complex associations.18,27 A good illustration of this is the complex [Ru(IP)2(dppz)]2+ (IP = imidazole[4,5-f][1,10]phenanthroline; Figure 11.1n), which, with its bulkier aromatic ancillary ligands, binds to DNA with an affinity several times that of its parent complex [Ru(bpy)2(dppz)]2+.46 Increased hydrophobicity in the form of strategically placed methyl groups can also strengthen the DNA-complex interaction: [Ru(5,6-Me2phen)2(dppz)]2+ (5,6-Me2phen = 5,6dimethyl-1,10-phenanthroline; Figure 11.1f) binds more strongly than does [Ru(phen)2(dppz)]2+.47 Ultimately, how well the ancillary ligands complement the shape of the groove in which they are binding (which itself depends on the nature of the target nucleic acid) may be the most significant factor governing binding.48 A detailed survey of DNA recognition by metallointercalators is beyond the scope of this review; however, an excellent review from the Barton laboratory has recently been published.49 Furthermore, rhodium(III)-based metallointercalators have been more extensively utilized for DNA recognition than ruthenium(II) complexes. However, the modification of the intercalating and ancillary ligands in [Ru(ancillary)2(intercalator)]2+ complexes has allowed some modulation of the binding selectivity to achieve specific DNA recognition. At the most basic level, ruthenium complexes can be used to distinguish between the A, B and Z-forms of DNA. For example, both the D and L enantiomers of [Ru(DIP)3]2+ bind to Z-form (left-handed) DNA, whereas only the D isomer binds to B-DNA;50 however, this was subsequently disputed by Norden and coworkers.11 Alternatively, [Ru(TMP)3]2+ (TMP = 3,4,7,8-tetramethyl-1,10-phenanthroline; Figure 11.1h) has been shown to specifically bind A-form DNA.51 More recently, it has been demonstrated that ruthenium-based metallointercalators can be used to target DNA mismatch sequences. These noncomplementary base pairs can occur naturally, or through the interaction of genotoxic chemicals or ionizing radiation, and can lead to a range of diseases.52 Barton and coworkers have reported that [Ru(bpy)2(tactp)]2+ (tactp = 4,5,9,18tetraazachrysenol(9,10-b)-triphenylene; Figure 11.1u) preferentially binds a CC mismatch.49 The tactp intercalating ligand combines the DNA light-switch effect of dppz-based complexes with the chrysi (5,6-chrysene quinone diimine; Figure 11.1v) ligand that is selective for mismatch sites. The sterically bulky chrysi ligand is too wide to intercalate at matched DNA sites, but is suited to thermodynamically destabilized sites.49

11.3 Dinuclear Complexes The utility of mononuclear metal complexes as probes for structural recognition in polynucleotides is limited by their relatively small size, as a typical mononuclear

Dinuclear Complexes

325

complex may, at best, span only four to six base pairs.53 Additionally, mononuclear complexes generally possess low binding affinities (typically ∼104–105 M−1, with the exception of strong intercalators like the dppz complexes) and consequently they are displaced from nucleic acids at relatively low ionic strengths, limiting their application in environments encountered in vivo.54 To mimic the selectivity of nucleic-acid binding proteins, larger species (spanning 10–16 base pairs) will be required: di-, tri- and oligonuclear metal complexes not only have a larger span, but they also possess a greater stereochemical diversity, allowing them to more effectively probe the shape- and structure-recognition mechanisms of nucleic acids than their mononuclear counterparts. Furthermore, dinuclear species possess a larger cationic charge; they may potentially have greater nucleic-acid binding affinities than mononuclear complexes; the presence of two metal centres amplifies any possible chiral discrimination effects; and the larger size of the complex results in considerably slower DNA dissociation rates (a property that is advantageous to both the biological activity and NMR studies of such complexes).55 The structures of bridging ligands discussed in this chapter are shown in Figure 11.2. The potential appeal of a dinuclear binding species can be exemplified by an early study of bis-intercalating organic molecules in which two intercalating chromophores (such as the ethidium cation) were joined via a flexible polyamine/alkane linker.56 Such dyes bound double-stranded DNA with an affinity several orders of magnitude higher than their monomeric analogues. As an example of a DNA-binding dinuclear metal complex, Aldrich-Wright et al. have investigated a similar flexible linker dimer, [{Ru(dpq)2}2(m-phen-x–SOS– x-phen)]4+ (SOS = 2-mercaptoethyl ether; the attachment position of the linker, x, is 4 or 5; dpq = dipyrido[3,2-d:2′,3′-f] quinoxaline; see Figure 11.3), featuring intercalating ancillary (terminal) ligands on each of the metal centres.53 The complex was found to have a binding affinity of approximately 6 × 107 M−1, far in excess of its mononuclear counterpart [Ru(dpq)2(phen)]2+ (K = 5.4 × 104 M−1).57 Furthermore, the complex demonstrates some degree of specificity for purine-purine sequences and has the potential for hydrogen-bonding interactions with the S–O–S bridge. There have also been a number of reports of complexes that bind via an intercalating bridging ligand. An early example of this was the 2,3-bis(2-pyridyl)benzo [g]quinoxaline (dpb; Figure 11.2d) bridged species described by Carlson et al.58 It was found that [{Ru(NH3)4}2(m-dpb)]4+ bound with an affinity similar to that of a mononuclear dppz-based intercalator, whereas [{Ru(bpy)2}2(m-dpb)]4+ bound rather weakly. This discrimination between the two complexes was attributed to the increased steric bulk of the bpy ancillary ligands interfering with complete insertion of the intercalating dpb moiety in the latter case and potential affinity-enhancing hydrogen bonding from the NH3 ligands in the former. Further studies on complexes in which the bridging ligand was extended – and potentially intercalating − revealed very interesting binding dynamics. The stereoisomers of the complex [{m-dppz(11,11′)dppz}(pp)4Ru2]4+ (dppz(11,11′)dppz = 11,11′bis(dipyrido[3,2-a:2′,3′-c]phenazine; Figure 11.2g; pp = bpy or phen) initially assumed a metastable major-groove-bound state at slightly different orientations for the various stereoisomers before threading through the double helix so that there was one Ru(phen)2 moiety in each groove, with the bridge intercalated.59

326

Groove-Binding Ruthenium(II) Complexes N

N

N

N

(f) pztp N

N

(a) bpm

N N N

N

N N

N

N

N

N

N

(b) HAT

N

N N

N

N

N

N

N

N

N

N

(g) dppz(11,11')dppz

N

(c) dppm N

N

N

N

CN

N N

O HN

N

(h) C4(cpdppz)2

N

(d) dpb

N

HN N

N

N

N

O

N

N (e) 2,3-dpp

N

Figure 11.2 Bridging ligands

CN

Dinuclear Complexes

327

N

N

NH

HN N

N

(i) bipp N

N N

N

H3C

N

N

N

N

N

N

N H2C

H3C

(j) tpphz

N

N

n

(k) bbn

N

Figure 11.2 Continued

4+ S

N (dpq)2Ru

N

S

N

O N

Ru(dpq)2

Figure 11.3 Representation of [{Ru(dpq)2}2(m-phen-5–SOS–5-phen)]4+.53

The introduction of some greater degree of flexibility between the two bridging dppz moieties was found to result in more expedient threading. The complex [mC4(cpdppz)2(phen)4Ru2]4+ (C4(cpdppz)2 = N,N′-bis(12-cyano-12,13-dihydro-11H-8cyclopenta[b]dipyrido[3,2-h:2′,3′-j]phenazine-12-carbonyl)-1,4-diaminobutane; Figure 11.2h) threaded through the DNA helix such that the ancillary phen ligands sat in the minor groove while the flexible linker occupied the major groove.60,61 Due to the bis-intercalative nature of the bridging ligand, the complex bound with an affinity several orders of magnitude greater than the analogous mononuclear species [Ru(phen)2(dppz)]2+ (∼1010 M−1 versus 106 M−1).61 The threading process is believed to require extensive conformational changes to the DNA helix, including the breaking and re-forming of base pairs. This is supported by the observation that, as with

328

Groove-Binding Ruthenium(II) Complexes

the dppz(11,11′)dppz-bridged species, AT-threading was faster than that for GC-rich sequences due to the stronger base-pairing of the latter. Kelly and coworkers have compared the binding behaviour of the dinuclear complex [(bpy)2Ru{bbn}Ru(bpy)2]4+ (bbn: n = 5 {1,5-bis[4(4′-methyl-2,2′bipyridyl)]pentane; n = 7 {1,7-bis[4(4′-methyl-2,2′-bipyridyl)]heptane}; Figure 11.2k) with its mononuclear analogues [Ru(bpy)3]2+ and [Ru(bpy)2(Me2bpy)]2+ (Me2bpy = 4,4′-dimethyl-2,2′-bipyridine; Figure 11.1b) and found that the bimetallic species had a much higher binding affinity, was more efficient at photosensitising DNA strand breaks and its binding affinity was less sensitive to ionic strength.62,63 Additional investigations were undertaken with [(phen)2Ru{bbn}Ru(phen)2]4+ (where n = 5, 7, or 10).63 Again, the dinuclear complex was found to have a stronger binding affinity than its mononuclear counterpart, [Ru(phen)2(Me2bpy)]2+, and that the binding affinity was dependent upon the linker chain length. The most effective binding was observed at n = 7, somewhat shorter than the optimal chain length (n > 8) of the classical intercalators with polymethylene chains.56 This shorter chain length is consistent with partial intercalation of a phenanthroline ligand on each metal centre.54 Other intercalative bridging moieties used in the study of DNA-binding dinuclear ruthenium complexes include phenanthroline/imidazole derivatives 64,65 and porphyrins,66 each of which yielded only moderate binding affinities akin to those observed in mononuclear species which bind via intercalation. One particularly interesting species utilized the intercalative bridge bipp (bipp = 2,9-bis(2-imidazo[4,5f][1,10]phenanthroline)-1,10-phenanthroline; Figure 11.2i) which features vacant chelating sites well suited to the binding of a third metal ion, specifically Cu2+. The emissive properties of the complex were quenched upon binding of the copper ion, whereas the DNA-bound complex exhibited a large increase in emission. Subsequent addition of copper ions to the complex–DNA system had no effect because the chelating site was blocked upon intercalation.67,68 This is in contrast with the socalled ‘molecular nut and bolt’ system of [Ru(bpy)2(tpphz)]2+ (tpphz = tetrapyrido[3,2a:2′,3′-c:3′′,2′′-h:2′′,3′′-j]phenazine; Figure 11.2j), which possesses a similar chelating site on the tpphz ligand. Upon intercalation with DNA, the chelating site projects out of the opposite side of the helix where it can coordinate a copper ion, quenching the emission of the complex and locking it in place.69 Many multinuclear species incorporating mixed-metal polypyridyl systems have shown considerable promise as photoactivated drugs: trinuclear Ru–Rh species (essentially two Ru(L)2 moieties linked to a RhCl2 core via 2,3-dpp bridges (2,3-dpp = 2,3-bis(2-pyridyl)pyrazine; Figure 11.2e)), for instance, have demonstrated visible light-induced photocleavage of DNA.70 This has been attributed to a Ru → Rh metal-to-metal charge transfer excited state and is dependent upon the nature of the bridging ligand (the analogous complex with 2,2′-bipyrimidine bridges (bpm; Figure 11.2a) was inactive).71 Ru/Pt mixed species, bridged by 2,3-dpp, dpq and dpb ligands, have been found to bind DNA covalently via the square-planar platinum moiety.72,73 Several nonintercalative, semi-rigid dinuclear complexes have also been investigated. Complexes bridged by the asymmetric phenanthroline derivative pztp ((3pyrazin-2-yl)-as-triazino[5,6-f]-1,10-phenanthroline; Figure 11.2f) have been found to bind via electrostatic/groove binding interactions, whereas the mononuclear

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equivalent intercalates.74 Furthermore, the groove complementarity necessary for intercalation means that the mononuclear complex exhibits enantioselectivity, whereas the (relatively) loosely associating dinuclear species does not. Jiang and coworkers have surveyed a number of dinuclear complexes featuring bridges derived from bpy.75–78 They all bind via a nonintercalative mode with modest binding affinities, however in a number of cases they exhibit intriguing enantiopreferences in which the L enantiomer appears to be the stronger binding.75,78 11.3.1 Stereoisomers and Their Separation A critical issue in the investigation of the interaction of the dinuclear species with DNA has been the ability to control the stereochemistry of the component metal centres. This has been achieved by a combination of techniques – stereoselective synthesis and chromatography.79,80 The stereoselective synthesis has utilized the chiral mononuclear precursors D/L-[Ru(pp)2(py)2]2+ 81 and D/L-[Ru(pp)2(CO)2]2+/ [Ru(pp)(pp′)(CO)2]2+ 82 (pp and pp′ are bidentate polypyridyl ligands; pp ≠ pp′), which can undergo substitution under conditions which allow the retention of the chiral integrity of the metal centre. The chromatographic technique has involved cation exchange, in which the counteranion has been chosen so that it differentially associates with the stereoisomers to be separated.83–85 The technique has allowed the routine separation of diastereoisomers,86–91 the resolution of enantiomers,86,89,90 the separation of geometric isomers89,92 and the separation of chiral helical forms.93 The combination of the stereoselective and chromatographic techniques has been used for the separation of stereoisomers of dinuclear and trinuclear species.92,94,95 11.3.2 Interaction of Dinuclear Complexes with Duplex DNA 1

H NMR experiments with the DL, DD and LL stereoisomers of [{Ru(Me2bpy)2}2(mbpm)]4+ (see Figure 11.4) demonstrated that the ruthenium(II) complexes bound duplex DNA in the minor groove, but with relatively weak affinity (K ≈ 103 M−1).90 Given the relative dimensions of the metal complex and the DNA grooves, it was somewhat surprising that the ruthenium complex bound in the minor groove: the

M

M

meso {Λ∆}

M

M

{ΛΛ}

M

rac

M

{∆∆}

Figure 11.4 A schematic representation of the three stereoisomers of a symmetric dinuclear ruthenium complex

330

Groove-Binding Ruthenium(II) Complexes

ruthenium complex is approximately 16 Å long and 8.0 Å wide, while for canonical form B-type DNA the width of the major and minor grooves are 11.6 Å and 6.0 Å, respectively. Although the ruthenium complexes bound duplex DNA weakly, the NMR results demonstrated that the DD- and LL-enantiomers interacted differently with the dodecanucleotides used in the study, with the DD-enantiomer having a slightly higher affinity. Mononuclear polypyridyl ruthenium(II) complexes that bind DNA nonintercalatively generally associate in the minor groove with a strong preference for A/T rich sequences.96 Distinctively, the stereoisomers of [{Ru(Me2bpy)2}2(m-bpm)]4+ were shown to bind d(CAATCCGGATTG)2 at the central CCGG sequence and at the CA/GT terminal residues, rather than at the expected A/T rich regions. As the minor groove is particularly narrow at A/T rich sequences, it is probable that the relatively bulky ruthenium complex could not be easily accommodated at the AAT/TTA sites. Consequently, it was proposed that the ruthenium complex binds at the central CCGG site, where the minor groove is more open, and at the terminal CA/GT site where the minor groove would also be considerably widened due to the fraying of the terminal base pair. 11.3.3 Interaction of Dinuclear Complexes with DNA Bulge Sequences As the bulky ruthenium complexes only bound DNA where the minor groove was relatively wide, it was proposed that they would bind DNA secondary structures that contain a more open or flexible groove with significantly higher affinity than duplex DNA. This proposal was consistent with the results from Kirsch-De Mesmaeker, Keene and coworkers,97 who showed that [{Ru(phen)2}2(m-HAT)]4+ (HAT = 1,4,5,8,9,12-hexaazatriphenylene; Figure 11.2b) binds partially-denatured DNA significantly more strongly than duplex DNA. One example of an open DNA structure is a sequence that contains a so-called ‘bulge’ – the inclusion of one or more bases on one strand that have no base(s) on the complementary strand with which to form a base pair (see Figure 11.5). DNA bulge sites can be created during recombination between imperfectly homologous sequences, and are thought to play an important role in frame-shift mutagenesis and be specific recognition sites for some DNA binding proteins. Consequently, there is considerable interest in developing small molecules that target bulge sites. NMR experiments demonstrated that DD-[{Ru(Me2bpy)2}2(m-bpm)]4+ bound an oligonucleotide containing a single adenine bulge significantly more strongly than the corresponding control, nonbulged, oligonucleotide.98 Furthermore, NOESY experiments indicated that the ruthenium complex bound specifically at the bulge site in the self-complementary tridecanucleotide d(CCGAGAATTCCGG)2 (where the A in bold represents the adenine bulge). Simple binding models, consistent with

Figure 11.5 A structural tridecanucleotide

representation

of

the

adenine

bulge

(A4)

containing

Dinuclear Complexes

331

Figure 11.6 Molecular model of DD-[{Ru(bpy)2}(m-bpm){Ru(Me2bpy)2}]4+ bound to d(CCGAGAATTCCGG)2.98

the NOE data, showed that the adenine bulge site was intrahelical and base-stacked with the adjacent guanine residues (see Figure 11.6). The binding model also indicated that the DNA minor groove had significantly widened at the bulge site to accommodate the ruthenium complex. As the minor groove at the bulge site in the free tridecanucleotide was not significantly wider than ‘canonical’ DNA, it was concluded that the bulge site introduces an increased flexibility into the local DNA structure that allows the specific metal complex binding. Interestingly, while the DD-enantiomer of [{Ru(Me2bpy)2}2(m-bpm)]4+ bound relatively strongly at the bulge site of d(CCGAGAATTCCGG)2, the corresponding LL-isomer did not bind at the bulge site, with weak binding to the terminal nucleotide residues being observed. The study provided a rare example of total enantioselectivity in the binding of an inert transition metal complex with DNA.99,100 The meso-(DL)-diastereoisomer bound with an affinity intermediate between the DDand LL-enantiomers. While it had been demonstrated that dinuclear ruthenium complexes could target single base bulges in DNA, it was not known if these metal complexes have potential for the larger bulge sites (three or more bases) often found in DNA and

332

Groove-Binding Ruthenium(II) Complexes

RNA. Molecular modelling suggested that the insertion of a three or more base bulge into duplex DNA should induce a bend towards the major groove that subsequently decreases the amount of helical twist at the bulge site.101 Consequently, the minor groove should be significantly straighter at the bulge site than in standard canonical form DNA, and therefore, a potentially good binding site for the rigid dinuclear ruthenium complexes. DD-[{Ru(phen)2}2(m-dppm)]4+ (dppm = 4,6-bis(2-pyridyl)pyrimidine; Figure 11.2c) was shown to bind at the A3-bulge site of the nonself-complementary dodecanucleotide duplex d(GCATCGAAAGCTACG)•d(CGTAGCCGATGC) with high affinity (K = 3 × 105 M−1) and selectivity.102 Interestingly, the NMR results suggested that at least one of the phen ligands is bound by partial intercalation. Upfield shifts of similar magnitude were observed for the phen resonances in a study of the binding of the well-known intercalating agent [Pt(en)(phen)]2+ (en = ethylenediamine) to a hexanucleotide.103 While the minor groove is no wider at the A3-bulge site compared to the normal duplex regions, molecular models showed that the smaller helical twist coupled with the 50 ° bend towards the major groove results in the minor groove being straighter at the A3-bulge sequence. This allows the bridging dppm ligand to follow the groove, and hence, fit more fully within the groove at the bulge site. Furthermore, possible steric clashes with the oligonucleotide backbone are also reduced for the two phenanthroline ligands that are almost parallel with the bridging dppm ligand. The remaining two phenanthroline ligands are then ideally placed to be inserted deeply within the duplex structure, as shown in Figure 11.7.

11.3.4 Interaction of Dinuclear Complexes with DNA Hairpin Sequences DNA hairpin sequences, or stem-loop structures – shown in Figure 11.8 – are secondary structural motifs known to arise naturally in base sequences possessing self-complementarity. Although not as common as in RNA, DNA hairpins and cruciforms (two opposed hairpins) are believed to play a role in a variety of fundamental biological processes, including the control of gene expression and mutagenic events.104–109 Many potential hairpin or cruciform structures have been identified in genomic DNA in regions associated with the regulation of transcription.105,106 Additionally, a number of DNA-binding proteins have been found to preferentially target such sequences.110–113 Conversely, a number of neurological diseases have been attributed to the disadvantageous formation of hairpin structures during the replication of trinucleotide repeat sequences.114,115 Hence, there is also significant interest in the design of small molecules that specifically target hairpin structures. The complexes meso-[{Ru(phen)2}2(m-HAT)]4+ and meso-[{Ru(4,7-Me2phen)2}2 (m-HAT)]4+ (4,7-Me2phen = 4,7-dimethyl-1,10-phenanthroline; Figure 11.1g) were shown to bind an icosanucleotide that contains a seven-base-pair stem section and a six-base loop region with greater affinity than the corresponding four-base loop octadecanucleotide and three control duplex dodecanucleotides.116 Based on the results from one- and two-dimensional NMR spectra, models of meso-[{Ru(phen)2}2 (m-HAT)]4+ and meso-[{Ru(4,7-Me2phen)2}2(m-HAT)]4+ bound to the six-base hairpin were produced. Each complex bound on the minor-groove side of the

Dinuclear Complexes

333

Figure 11.7 Two alternate orientations of an energy-minimized model of DD-[{Ru(phen)2}2(mdppm)]4+ bound at the bulge site of the A3-bulge oligonucleotide.102 The dinuclear ruthenium complex is coloured black

Figure 11.8 Schematic representation of a six-base hairpin loop sequence

334

Groove-Binding Ruthenium(II) Complexes

icosanucleotide, with the HAT ligand positioned at the stem-loop interface with one set of terminal phenanthroline ligands projecting into the groove of the stem, and the other projecting into the loop region. The groove at the stem-loop interface is wider, or more flexible, than the duplex stem of the icosanucleotide. It was suggested that the binding was stabilized by energetically favourable van der Waals and hydrophobic interactions with the bulky dinuclear complexes. Of the two metal complexes, meso-[{Ru(4,7-Me2phen)2}2(m-HAT)]4+ was found to bind the six-base hairpin-containing icosanucleotide more strongly; however, it also bound the control duplex sequences with significant affinity. Secretion of the hydrophobic methyl groups away in the minor groove, and the subsequent displacement of solvent, was proposed as the driving force behind this greater binding ability of the methylated species. Alternatively, while meso-[{Ru(phen)2}2(m-HAT)]4+ bound the six-base hairpin sequence less strongly than the corresponding methyl analogue, it showed greater selectivity in its binding when comparing the affinity to the hairpin sequence and control duplex structures. Metal complexes possessing phen or 4,7-Me2phen terminal ligands bound the icosanucleotide hairpin structure with greater affinity than the ruthenium complexes containing corresponding bpy and Me2bpy terminal ligands.116 Presumably, the larger surface areas of the phenanthroline ligand compared to bipyridine results in better van der Waals contacts with the walls of the minor groove, or more favourable hydrophobic interactions, or allows a semi-intercalative association with DNA bases, or a combination of all three. With regards to bridging ligands, the various studies demonstrated that ruthenium complexes containing a bpm bridge are generally selective for bulge sites, while a HAT bridging ligand is preferable for targeting hairpin structures. It was also noted that the meso diastereoisomer gives the strongest binding, particularly in those complexes possessing phenanthroline ligands. Simple computer models illustrated that the terminal ligands of a meso isomer are able to follow the contours of the minor groove, whereas the DD- and LL-enantiomeric forms of the complex encounter steric clashes at either end of the complex.116

11.3.5 Interaction of Dinuclear Complexes with RNA Sequences Although it has been clearly established that inert dinuclear ruthenium(II) complexes preferentially bind to nonduplex DNA structures, little is know about their interaction with RNA. Nonduplex structures occur in both DNA and RNA, but are much more prevalent in the latter. The formation of such structures is generally transient and/or disadvantageous in DNA. Alternatively, RNA exhibits a large diversity of nonduplex structures, which have an important role for recognition sites for proteins, RNA splicing and RNA folding, making them promising targets for potential diagnostic and therapeutic agents. Given the differences in the conformations of DNA and RNA, it is not possible to extrapolate the results obtained for DNA to RNA. Standard form DNA adopts a B-type helix, whereas RNA adopts an A-type structure. This results in significant differences in the groove dimensions – DNA contains a wide major groove and relatively narrow minor groove, whereas, RNA has a deep and very narrow major groove and a wide shallow minor groove (see Figure 11.9).117

Dinuclear Complexes

335

Figure 11.9 Comparative models of B- (left) and A-form (right) nucleic acids. The major and minor grooves are labelled

In order to make a direct comparison with the results obtained with DNA, the binding of the two enantiomers of [{Ru(Me2bpy)2}2(m-bpm)]4+ to the RNA analogue of the adenine bulge-containing DNA sequence d(CCGAGAATTCCGG)2, and the corresponding bulge-free control RNA dodecanucleotide, were examined.118 Both enantiomers bound the control RNA duplex weakly (K = 1 × 103 M−1), but bound the r(CCGAGAAUUCCGG)2 structure with considerably greater affinity (K = 6 × 104 M−1). From NOESY spectra, it was established that the ruthenium complex did selectively bind at the bulge site. A binding model that was consistent with all the NMR was obtained that demonstrated that DD-[{Ru(Me2bpy)2}2(mbpm)]4+ could effectively associate at the bulge site in the RNA minor groove. It was proposed that the insertion of the bulge into the tridecanucleotide formed a metal complex binding pocket by partially unwinding the duplex at the unpaired adenine. This perturbation to the RNA duplex caused a reduction in the curvature of the minor groove that may have aided the binding of the ruthenium complex. Alternatively, the minor groove structure at the bulge site may be more flexible, due to the reduced thermodynamic stability produced by the unpaired adenine, thereby allowing a better induced-fit binding site. The results of the study demonstrated that inert dinuclear ruthenium complexes have excellent potential as probes for nonduplex RNA structures, in a similar manner to that previously established for DNA. 11.3.6 Dinuclear Ruthenium Complexes with Flexible Ligands Although the dinuclear ruthenium complexes that are bridged by rigid linking ligands, such as bpm and HAT, exhibit relatively strong binding to nonduplex DNA

336

Groove-Binding Ruthenium(II) Complexes

and RNA structures, it is generally only possible for one metal centre to bind deeply within the minor groove. The rigid ruthenium complex cannot follow the curvature of the groove in nonduplex structures that are not dramatically altered from the standard duplex, and hence the second metal centre must project out of the groove. It was proposed that dinuclear ruthenium complexes linked by a flexible chain could overcome this limitation, and potentially allow both metal centres to bind optimally in the minor groove. The stereoisomers of a series of dinuclear ruthenium(II) complexes [{Ru(phen)2}2(m-bbn)]4+ with flexible bridging ligands (bbn; Figure 11.2k) – bb2 (1,2bis[4(4′-methyl-2,2′-bipyridyl)]ethane), bb5 (1,5-bis[4(4′-methyl-2,2′-bipyridyl)]pentane), bb7 (1,7-bis[4(4′-methyl-2,2′-bipyridyl)]heptane) and bb10 (1,10-bis[4(4′-methyl2,2′-bipyridyl)]decane} – were synthesized and their binding to the single adenine bulge-containing tridecanucleotide, d(CCGAGAATTCCGG)2, studied.119 Based on the results from a fluorescent DAPI-displacement assay (DAPI = 4′,6-diamidino-2phenylindole; see Section 11.3.7), equilibrium dialysis and affinity binding chromatography experiments it was determined that the DD-enantiomer of the bb7-bridged complex bound the bulge-containing oligonucleotide with the highest affinity. NMR experiments demonstrated that the DD-bb7-bridged complex still selectively targeted the bulge site, while molecular modelling showed the ability of the flexibly linked complex to follow the curvature of the DNA minor groove, as shown in Figure 11.10. It was concluded that flexibly linked dinuclear complexes will still exhibit the selectivity for nonduplex DNA sites observed for the rigid complexes,

Figure 11.10 Comparison of the binding of DD-[{Ru(phen)2}2(m-bb7)]4+(left) and DD[{Ru(phen)2}2(m-bpm)]4+ (right) to the adenine-bulged tridecanucleotide duplex. The complexes are coloured black, and the arrows indicate the location of the second of the ruthenium metals centres (after the first one has positioned itself in the bulge region) showing how in the flexibly linked complex the linker follows the curve in the groove119

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337

but they should be able to bind with higher affinity. No conclusive reason was proposed for why DD-[{Ru(phen)2}2(m-bb7)]4+ bound more strongly than the corresponding complexes containing two, five or ten methylene groups. It was considered likely that the bb2-bridged complex contains insufficient degrees of freedom to bind tightly, whereas the preference of the bb7-bridged complex over the bb5- and bb-10 bridged complexes may rely on the differential binding potential of the DNA sequence where the second ruthenium centre binds. 11.3.7 Fluorescent DAPI-Displacement Assay While NMR spectroscopy provides the most detailed information on the nature of the binding of the ruthenium complexes with any of the DNA or RNA structures, it is a relatively time-consuming and expensive technique (in terms of the cost of the oligonucleotides). A suitable method is required to screen the relative binding affinity of a large number of ruthenium complexes, in terms of variable ligands and stereoisomers, against a range of DNA and RNA structures. Boger et al.120 developed a now widely used fluorescent intercalator displacement (FID) assay to screen organic compounds that bind, by intercalation or groove association, with oligonucleotides. The assay is based upon the loss of fluorescence resulting from the displacement of an intercalating dye such as ethidium bromide (EthBr) or thiazole orange (TO) from a DNA sequence by the molecule of interest. While the assay appears to be widely applicable, some anomalies were observed with dinuclear ruthenium complexes between the results obtained with the FID method and observations on the same oligonucleotide–metal complex combinations using other techniques. As the disparity may have been due to differences in binding modes between the intercalating dyes and the minor groove-binding metal complexes, a modified fluorescence assay was proposed that used the minor groovebinding compound DAPI. It was suggested that there may be a direct displacement of the fluorescent DAPI dye by the minor groove-binding ruthenium complexes.121 In a systematic evaluation of the DNA-binding affinities of a range of dinuclear ruthenium complexes with various oligonucleotides, some notable inconsistencies were observed between the results of the FID fluorescence assay using an intercalating dye, compared with DAPI. The metal complex-oligonucleotide combinations that gave inconsistent results in the fluorescent displacement assays were further examined by other techniques, including NMR, affinity chromatography and equilibrium dialysis. The results from the DAPI assay were totally consistent with the relative metal complex-oligonucleotide affinities determined by the other techniques. It was concluded that the difference in the binding mode between the intercalating dye and the groove-binding ruthenium complexes were responsible for the discrepancies.121 11.3.8 Affinity Chromatography The interactions of DNA with metal complexes may be utilized in the ‘reverse’ sense to develop a remarkably efficient affinity chromatographic technique for the

338

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separation of stereoisomers of dinuclear species.122 The attachment of selected biotinylated oligonucleotides to a stationary phase comprising streptavidin immobilized on Sepharose has allowed separation over very short distances (2–5 cm) of mixtures of different complexes, the separation of geometric isomers and diastereoisomers of the same complex, and resolution of chiral forms – separations which have only been achieved by the conventional cation-exchange column techniques with an effective column length in excess of 30 metres!

11.4 Potential Biological Significance While it has been clearly demonstrated that dinuclear ruthenium complexes have considerable potential as probes for nonduplex DNA and RNA, this potential will only be realized if: (1) the metal complexes can be functionalized so that they selectively bind specific nonduplex structures that are biologically significant; and/or (2) the metal complexes can be transported into human cells, and thereby elicit some biological response, e.g. they are cytotoxic. Given the great diversity of organic ligands that can be readily synthesized, it is probable that once specific nonduplex sites are identified, hydrogen bond donors and acceptors and/or van der Waals interaction recognition groups can be incorporated into the metal complex. An excellent example of this type of approach was reported by Barton and coworkers, who specifically designed the DNA-intercalating complex D-a-[Rh{(R,R)-Me2trien}(phi)]3+ ((R,R)-Me2trien = 2R,9R-diamino-4,7diazadecane; Figure 11.1m) to bind to the sequence 5′-TGCA-3′.123 The intercalating ligand phi locks the rhodium complex into the DNA duplex. Additional recognition elements, hydrogen bonding NH2 groups and methyl groups that can make van der Waals contacts with the TMe groups, were then incorporated into the metal complex in the correct three-dimensional position to interact with their DNA counterpart.123 Although there has been one recent study,124 little is known about the cellular transport and cytotoxicity of dinuclear ruthenium complexes. However, the biological properties of a range of mononuclear ruthenium complexes have been reported. Puckett and Barton125 examined the cellular transport of a series of DNA-intercalating ruthenium complexes, and demonstrated that those with greater lipophilicity exhibit highest uptake, while charge and size were not particularly important. Furthermore, Liu et al.126 reported that [Ru(phen)2(PIP)]2+ (where PIP = R-phenylimidazo[4,5-f] [1,10]phenanthroline; Figure 11.1t), which binds DNA by intercalation, exhibited substantial cytotoxicity in a range of cell lines. Given that the cytotoxicity of many organic compounds is due to their ability to intercalate into the DNA helix, it is not surprising that ruthenium complexes that bind DNA in a similar mode also exhibit reasonable cytotoxicity. Similarly, examples of DNA groove-binding ruthenium complexes that are cytotoxic have been reported. Ma et al.127 demonstrated that although [Ru(tBu2bpy)2(2-appt)]2+ (tBu2bpy = 4,4′bis(tert-butyl)-2,2′-bipyridine; Figure 11.1c; 2-appt = 2-amino-4-phenylamino-6-(2-

Abbreviations of Ligands

339

pyridyl)-1,3,5-triazine; Figure 11.1r) bound DNA with only modest affinity, it also exhibited moderate cytotoxicity in several human cancer cell lines. Additionally, Spillane et al.128 reported that the DNA minor groove binding metal complex [Ru(bpy)2(bbtb)]2+ (bbtb = 4,4′-bis(benzothiazol-2-yl)-2,2′-bipyridine; Figure 11.1s) showed significant cytotoxicity in a range of human cancer cell lines. Due to the success (and associated clinical problems) of cisplatin as an anticancer agent, there has been intense interest in transition-metal-based compounds that bind DNA. Through the early studies by the research groups of Barton and Nordén, ruthenium complexes that can bind DNA and RNA by intercalation are now well-established probes of nucleic acids. Over the last five or so years, dinuclear ruthenium complexes that can only bind nucleic acids through association in the minor groove have also become recognized as valuable probes for nonduplex nucleic acid structures. As nonduplex nucleic acid structures have been shown to be important in the regulation of gene expression, di- and oligonuclear ruthenium complexes may have significant potential as therapeutic agents for a range of diseases. Furthermore, given the vast amount of classical inorganic chemistry research that has involved ruthenium(II) complexes, it is probable that this class of nucleic acidbinding metal complex can be readily tuned to achieve the desired biological outcomes.

Abbreviations of Ligands 2-appt bbn

bbtb bipp bpm bpy t Bu2bpy C4(cpdppz)2

chrysi DIP dpb 2,3-dpp dppm dppx dppz dppz(11,11′)dppz

2-amino-4-phenylamino-6-(2-pyridyl)-1,3,5-triazine, Figure 11.1r bb2 {1,2-bis[4(4′-methyl-2,2′-bipyridyl)]ethane}, bb5 {1,5bis[4(4′-methyl-2,2′-bipyridyl)]pentane}, bb7 {1,7-bis[4(4′methyl-2,2′-bipyridyl)]heptane}, and bb10 {1,10-bis[4(4′-methyl-2,2′-bipyridyl)]decane}, Figure 11.2k 4,4′-bis(benzothiazol-2-yl)-2,2′-bipyridine, Figure 11.1s 2,9-bis(2-imidazo[4,5-f][1,10]phenanthroline)-1,10phenanthroline, Figure 11.2i 2,2′-bipyrimidine, Figure 11.2a 2,2′-bipyridine, Figure 11.1a 4,4′-bis(tert-butyl)-2,2′-bipyridine, Figure 11.1c N,N′-bis(12-cyano-12,13-dihydro-11H-8-cyclopenta[b]dipyrido[ 3,2-h:2′,3′-j]phenazine-12-carbonyl)-1,4-diaminobutane, Figure 11.2h 5,6-chrysene quinone diimine, Figure 11.1v 4,7-diphenyl-1,10-phenanthroline, Figure 11.1o 2,3-bis(2-pyridyl)benzo[g]quinoxaline, Figure 11.2d 2,3-bis(2-pyridyl)pyrazine, Figure 11.2e 4,6-bis(2-pyridyl)pyrimidine, Figure 11.2c 7,8-dimethyldipyridophenazine, Figure 11.1l dipyrido[3,2-a:2′,3′-c]phenazine, Figure 11.1j 11,11′-bis(dipyrido[3,2-a:2′,3′-c]phenazine, Figure 11.2g

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Groove-Binding Ruthenium(II) Complexes

dpq dpqC HAT IP Me2bpy 2,9-Me2phen 5,6-Me2phen 4,7-Me2phen (R,R)Me2trien phen phi picchxnMe2 PIP pztp tactp TMP tpphz

dipyrido[3,2-d:2′,3′-f]quinoxaline, Figure 11.1i dipyrido[3,2-a:2′,3′-c](6,7,8,9-tetrahydro)phenazine, Figure 11.1k 1,4,5,8,9,12-hexaazatriphenylene, Figure 11.2b imidazole[4,5-f][1,10]phenanthroline, Figure 11.1n 4,4′-dimethyl-2,2′-bipyridine, Figure 11.1b 2,9-dimethyl-1,10-phenanthroline, Figure 11.1e 5,6-dimethyl-1,10-phenanthroline, Figure 11.1f 4,7-dimethyl-1,10-phenanthroline, Figure 11.1g 2R,9R-diamino-4,7-diazadecane, Figure 11.1m 1,10-phenanthroline, Figure 11.1d phenanthrenequinone diimine, Figure 11.1p N,N′-dimethyl-N,N′-di(2-picolyl)-1,2-diaminocyclohexane, Figure 11.1q R-phenylimidazo[4,5-f][1,10]phenanthroline, Figure 11.1t 3-pyrazin-2-yl)-as-triazino[5,6-f]-1,10-phenanthroline, Figure 11.2f 4,5,9,18-tetraazachrysenol(9,10-b)-triphenylene, Figure 11.1u 3,4,7,8-tetramethyl-1,10-phenanthroline, Figure 11.1h tetrapyrido[3,2-a:2′,3′-c:3′′,2′′-h:2′′,3′′-j]phenazine, Figure 11.2j

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72. Milkevitch, M.; Storrie, H.; Brauns, E.; Brewer, K.J.; Shirley, B.W.; A new class of supramolecular, mixed-metal DNA-binding agents: the interaction of RuII,PtII and OsII,PtII bimetallic complexes with DNA; Inorg. Chem., 1997, 36, 4534–4538. 73. Williams, R.L.; Toft, H.N.; Winkel, B.; Brewer, K.J.; Synthesis, characterization, and DNA binding properties of a series of Ru, Pt mixed-metal complexes; Inorg. Chem., 2003, 42, 4394–4400. 74. Zou, X.-H.; Ye, B.-H.; Li, H.; Liu, J.-G.; Xiong, Y.; Ji, L.-N.; Mono- and bi-nuclear ruthenium(II) complexes containing a new asymmetric ligand 3-(pyrazin-2-yl)-as-triazino[5,6-f]1,10-phenanthroline: synthesis, characterization and DNA-binding properties; J. Chem. Soc., Dalton Trans., 1999, 1423–1428. 75. Jiang, C.-W.; Chao, H.; Hong, X.-L.; Li, H.; Mei, W.-J.; Ji, L.-N.; Enantiopreferential DNA-binding of a novel dinuclear complex [(bpy)2Ru(bdptb)Ru(bpy)2]4+; Inorg. Chem. Commun., 2003, 6, 773–775. 76. Jiang, C.-W.; Syntheses, characterization and DNA binding of mono- and diruthenium(II) complexes containing 2,2′-bis(1,2,4-triazino[5,6-f]phenanthren-3-yl)-4,4′-bipyridine and 2,2′-bipyridine ligands; Inorg. Chim. Acta, 2005, 358, 1231–1236. 77. Jiang, C.-W.; Chiral diruthenium(II) complexes containing a modified 4,4′-bipyridine bridging ligand: syntheses, characterization and DNA binding; Inorg. Chim. Acta, 2004, 357, 3403–3406. 78. Jiang, C.-W.; Dimeric 2,2-bipyridylruthenium(II) complexes containing 2,2-bis(1,2,4triazin-3-yl)-4,4-bipyridine-like bridging ligands: Syntheses, characterization and DNA binding; Eur. J. Inorg. Chem., 2004, 2277–2282. 79. Keene, F.R.; Stereochemistry and polymetallic ligand-bridged molecular assemblies; Coord. Chem. Rev., 1997, 166, 121–159. 80. Keene, F.R.; Isolation and characterisation of stereoisomers in di- and tri-nuclear complexes; Chem. Soc. Rev.; 1998, 27, 185–193. 81. Hua, X.; von Zelewsky, A.; Enantiomerically pure chiral RuII(L-L)2 building blocks for coordination compounds; Inorg. Chem.; 1995, 34, 5791–5797. 82. Rutherford, T.J.; Quagliotto, M.G.; Keene, F.R.; Chiral [Ru(pp)2(CO)2]2+ species (pp = bidentate polypyridyl ligand) and their use in the stereoselective synthesis of ligandbridge dinuclear complexes; Inorg. Chem., 1995, 34, 3857–3858. 83. Rutherford, T.J.; Pellegrini, P.A.; Aldrich-Wright, J.R.; Junk, P.C.; Keene, F.R.; Isolation of enantiomers of a range of tris(bidentate)ruthenium(II) species using chromatographic resolution and stereoretentive synthetic methods; Eur. J. Inorg. Chem., 1998, 1677–1688. 84. Fletcher, N.C.; Junk, P.C.; Reitsma, D.A.; Keene, F.R.; Chromatographic separation of stereoisomers of dinuclear ligand-bridged polypyridyl complexes of ruthenium(II); J. Chem. Soc., Dalton Trans., 1998, 133–138. 85. Fletcher, N.C.; Keene, F.R.; Anion interactions with (polypyridyl)ruthenium complexes, and their importance in the cation-exchange chromatographic separation of stereoisomers of dinuclear species; J. Chem. Soc., Dalton Trans., 1999, 683–689. 86. Kelso, L.S.; Reitsma, D.A.; Keene, F.R.; Spectral and electrochemical properties of the diastereoisomeric forms of azobis(2-pyridine)-bridged diruthenium species; Inorg. Chem., 1996, 35, 5144–5153. 87. Richardson, C.; Steel, P.J.; D’Alessandro, D.M.; Junk, P.C.; Keene, F.R.; Mono- and dinuclear complexes of the ligands 3,4-di(2-pyridyl)-1,2,5-oxadiazole and 3,4-di(2-pyridyl)1,2,5-thiadiazole; new bridges allowing unusually strong metal-metal interactions; J. Chem. Soc., Dalton Trans., 2002, 2775–2785. 88. Zampese, J.A.; Keene, F.R.; Steel, P.J.; Diastereoisomeric dinuclear ruthenium complexes of 2,5-di(2-pyridyl)thiazolo[5,4d]thiazole; Dalton Trans. 2004, 4124–4129. 89. Patterson, B.T.; Foley, F.M.; Richards, D.; Keene, F.R.; Synthetic and stereochemical studies of dinuclear ruthenium(II) with mixed terminal ligands; Dalton Trans., 2003, 709–714. 90. Foley, F.M.; Keene, F.R.; Collins, J.G.; The DNA binding of the DD-, DL- and LLstereoisomers of [{Ru(Me2bpy)2}2(m-bpm)]4+; J. Chem. Soc., Dalton Trans., 2001, 2968– 2974.

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91. D’Alessandro, D.M.; Keene, F.R.; Diastereoisomers as probes for solvent reorganizational effects on IVCT in dinuclear ruthenium complexes; Chem. Phys., 2006, 324, 8–25. 92. Patterson, B.T.; Keene, F.R.; Isolation of geometric isomers within diastereoisomers of dinuclear ligand-bridged complexes; Inorg. Chem., 1998, 37, 645–650. 93. Rapenne, G.; Patterson, B.T.; Sauvage, J.-P.; Keene, F.R.; Resolution, X-ray structure and absolute configuration of a double-stranded helical diiron(II) bis(terpyridine) complex; Chem. Commun., 1999, 1853–1854. 94. Rutherford, T.J.; Keene, F.R.; Stereoisomers in heterometallic (Ru2Os) and heteroleptic homometallic (RuRu′Ru′′) trinuclear complexes incorporating the bridging ligand HAT (1,4,5,8,9,12-hexaazatriphenylene); J. Chem. Soc., Dalton Trans., 1998, 1155–1162. 95. Rutherford, T.J.; Van Gijte, O.; Kirsch-De Mesmaeker, A.; Keene, F.R.; Stereoisomers of mono-, di- and tri-ruthenium(II) complexes containing the bridging ligand 1,4,5,8,9,12hexaazatriphenylene (HAT), and studies of their photophysical and redox properties; Inorg. Chem., 1997, 36, 4465–4474. 96. Aldrich-Wright, J.R.; Greguric, I.D.; Lau, C.H.Y.; Pellegrini, P.; Collins, J.G.; Transition metal complexes as probes for DNA; Recent Res. Devel. Inorg. Chem., 1998, 1, 13–36. 97. Broadkorb, A.; Kirsch-De Mesmaeker, A.; Rutherford, T.J.; Keene, F.R.; Stereoselective interaction and photo-electron transfer between mononucleotides or DNA and the stereoisomers of a dinuclear Ru(II) HAT complex; Eur. J. Inorg. Chem., 2001, 2151–2160. 98. Patterson, B.T.; Collins, J.G.; Foley, F.M.; Keene, F.R.; Dinuclear ruthenium(II) complexes as probes for DNA bulge sites; J. Chem. Soc., Dalton Trans., 2002, 4343–4350. 99. Smith, J.A.; Collins, J.G.; Patterson, B.T.; Keene, F.R.; Total enantioselectivity in the DNA binding of the dinuclear ruthenium(II) complex [{Ru(Me2bpy)2}2(m-bpm)]4+ {bpm = 2,2′bipyrimidine; Me2bpy = 4,4′-dimethyl-2,2′-bipyridine}; Dalton Trans., 2004, 1277– 1283. 100. Morgan, J.L.; Buck, D.P.; Turley, A.G.; Collins, J.G.; Keene, F.R.; Meso-[{Ru(phen)2}2(mbpm)]4+: a high-affinity DNA bulge probe {bpm = 2,2′-bipyrimidine; phen = 1,10-phenanthroline}; Inorg. Chim. Acta, 2006, 359, 888–898. 101. Rosen, M.A.; Live, D.; Patel, D.J.; Comparative NMR study of AN-bulge loops in DNA duplexes: intrahelical stacking of A, A-A, and A-A-A bulge loops; Biochemistry, 1992, 31, 4004–4014. 102. Morgan, J.L.; Buck, D.P.; Turley, A.G.; Collins, J.G.; Keene, F.R.; Selectivity at a three-base bulge site in the DNA binding of DD-[{Ru(phen)2}2(m-dppm)]4+ {dppm = 4,6-bis(2pyridyl)pyrimidine; phen = 1,10-phenanthroline}; J. Biol. Inorg. Chem., 2006, 11, 824–834. 103. Collins, J.G.; Rixon, R.M.; Aldrich-Wright, J.R.; Interaction of [Pt(en)(phen)]2+ and [Pt(en)(phi)]2+ with the hexanucleotide d(GTCGAC)2: Evidence for minor groove binding; Inorg. Chem., 2000, 39, 4377–4379. 104. Batey, R.T.; Rambo, R.P.; Doudna, J.A.; Tertiary motifs in RNA structure and folding; Angew. Chem. Int. Ed., 1999, 38, 2326–2343. 105. Varani, G.; Exceptionally stable nucleic acid hairpins; Annu. Rev. Biophys. Biomol. Struct., 1995, 24, 379–404. 106. Mizuuchi, K.; Mizuuchi, M.; Gellert, M.; Cruciform structures in palindromic DNA are favored by DNA supercoiling; J. Mol. Biol., 1982, 156, 229–243. 107. Dai, X.; Greizerstein, M.B.; Nadas-Chinni, K.; Rothman-Denes, L.B.; Supercoil-induced extrusion of a regulatory DNA hairpin; Proc. Natl. Acad. Sci. USA, 1997, 94, 2174–2179. 108. Spiro, C.; McMurray, C.T.; Switching of DNA secondary structure in proenkephalin transcriptional regulation; J. Biol. Chem., 1997, 272, 33145–33152. 109. Glucksmann, M.A.; Markiewicz, P.; Malone, C.; Rothman-Denes, L.B.; Specific sequences and a hairpin structure in the template strand are required for N4 virion RNA polymerase promoter recognition; Cell, 1992, 70, 491–500. 110. Zazopoulos, E.; Lalli, E.; Stocco, D.M.; Sassone-Corsi, P.; DNA binding and transcriptional repression by DAX-1 blocks steroidogenesis; Nature, 1997, 390, 311–315.

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111. Froelich-Ammon, S.J.; Gale, K.C.; Osheroff, N.; Site-specific cleavage of a DNA hairpin by topoisomerase II. DNA secondary structure as a determinant of enzyme recognition/cleavage; J. Biol. Chem., 1994, 269, 7719–7725. 112. Hao, Q.; Peumans, W.J.; van Damme, E.J.M.; Type-1 ribosome-inactivating protein from iris (Iris hollandica var. Professor Blaauw) binds specific genomic DNA fragments; Biochem. J., 2001, 357, 875-880. 113. Lilley, D.M.; The inverted repeat as a recognizable structural feature in supercoiled DNA molecules; Proc. Natl. Acad. Sci. USA, 1980, 77, 6468–6472. 114. Sutherland, G.R.; Richards, R.I.; Simple tandem DNA repeats and human genetic disease; Proc. Natl. Acad. Sci. USA, 1995, 92, 3636–3641. 115. Pearson, C.E.; Edamura, K.N.; Cleary, J.D.; Repeat instability: mechanisms of dynamic mutations; Nat. Rev. Genet., 2005, 6, 729–742. 116. Smith, J.A.; Morgan, J.L.; Turley, A.G.; Collins, J.G.; Keene, F.R.; Meso-[{Ru(phen)2}2(mHAT)]4+: a high-affinity DNA hairpin probe (HAT = 1,4,5,8,9,12-hexaazatriphenylene; phen = 1,10-phenantroline); Dalton Trans., 2006, 3179–3187. 117. Neidle, S.; DNA Structure and Recognition, IRL Press, Oxford, 1994. 118. Spillane, C.B.; Smith, J.A.; Buck, D.P.; Collins, J.G.; Keene, F.R.; Dinuclear ruthenium(II) complexes as potential probes for RNA bulge sites; Dalton Trans., 2007, 5290–5296. 119. Morgan, J.L.; Spillane, C.B.; Smith, J.A.; Buck, D.P.; Collins, J.G.; Keene, F.R.; Dinuclear ruthenium(II) complexes with flexible bridges as DNA bulge-selective probes; Dalton Trans. 2007, 4333–4342. 120. Boger, D.L.; Fink, B.E.; Brunette, S.R.; Tse, W.C.; Hedrick, M.P.; A simple, high-resolution method for establishing DNA binding affinity and sequence selectivity; J. Am. Chem. Soc., 2001, 123, 5878–5891. 121. Spillane, C.B.; Smith, J.A.; Morgan, J.L.; Keene, F.R.; DNA affinity binding studies using a fluorescent dye displacement technique – the dichotomy of the binding site; Chem., 2007, 12, 819–824. 122. Smith, J.A.; Keene, F.R.; Separation of stereoisomers of dinuclear metal complexes by binding affinity chromatography using non-duplex DNA; Chem. Commun., 2006, 2583–2585. 123. Hudson, B.P.; Dupureur, C.M.; Barton, J.K.; 1H NMR structural evidence for the sequence-specific design of an intercalator: D-a-[Rh[(R,R)-Me2trien]phi]3+ bound to d(GAGTGCACTC)2; J. Am. Chem. Soc., 1995, 117, 9379–9380. 124. McDonnell, U.; Kerchoffs, J.M.C.A.; Castineiras, R.P.M.; Hicks, M.R.; Hotz, A.C.G.; Hannon, M.J.; Rodger, A.; Synthesis and cytotoxicity of dinuclear complexes containing ruthenium(II) bipyridyl units linked by a bis(pyridylimine) ligand; Dalton Trans., 2008, 667–675. 125. Puckett, C.A.; Barton, J.K.; Methods to explore cellular uptake of ruthenium complexes; J. Am. Chem. Soc., 2007, 129, 46–47. 126. Liu, J.; Zheng, W.; Shi, S.; Tan, C.; Chen, J.; Zheng. K.; Ji, L.; Synthesis, antitumor activity and structure–activity relationships of a series of Ru(II) complexes; J. Inorg. Biochem., 2008, 102, 193–202. 127. Ma, D.-L.; Che, C.-M.; Siu, F.-M.; Yang, M.; Wong, K.-Y.; DNA binding and cytotoxicity of ruthenium(II) and rhenium(I) complexes of 2-amino-4-phenylamino-6-(2-pyridyl)1,3,5-triazine; Inorg. Chem., 2007, 46, 740–749. 128. Spillane, C.B.; Fletcher, N.C.; Roundtree, S.; van den Berg, H.; Chanduloy, S.; Morgan, J.L.; Keene, F.R.; Benzothiazole bipyridine complexes of ruthenium(II) with cytotoxic activity; J. Biol. Inorg. Chem., 2007, 12, 797–807.

12 DNA Recognition and Binding by Peptide–Metal Complex Conjugates Alexandra Myari and Nick Hadjiliadis

12.1

Introduction

The design and synthesis of chemical agents capable of binding specifically to DNA have gained great research interest in recent years.1 The discovery of novel sequence specificities, beyond the sequences recognized by the known natural restriction endonucleases, may possibly offer extended possibilities in DNA manipulation.2 Such specific DNA-binding agents could find several applications in molecular biology (sequencing, cloning) and clinical applications (gene regulation, gene therapy, chemotherapy).2 X-ray crystallography3,4 has revealed many molecular details regarding protein– DNA recognition. These involve the formation of a hydrogen-bonding network between peptide donor groups (coming from the peptide backbone and amino acid side chains) and the edges of the bases, consisting of what is called direct readout. In addition, formation of specific complexes between the protein and DNA is often followed by conformational changes of both protein and DNA molecules as well as release of water molecules part of the DNA hydration spine.3–5 Peptides are low molecular weight DNA-binding agents that can effectively mimic the bulky proteins. Peptides have the same donor groups utilized by proteins in the process of DNA recognition. They are also easily synthesized and their amino-acid composition and chirality can be tuned in order to achieve the desired DNA-binding properties. Binding of peptides has been thoroughly investigated and Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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sequence selectivity has been demonstrated in many cases.6 However, short peptides lack the DNA-binding affinity of the native proteins. This is why dimeric peptides have been used for such studies, instead.7 Peptides can become powerful tools in our hands if a specific protein–DNA recognition code is determined. There is a big debate between different research groups as to whether a ‘DNA recognition code’ exists.8 As many structures of DNA–transcription factor complexes became known in atomic detail, it was recognized that some DNA-binding proteins share the same structural framework, such as helix-turn-helix, zinc finger and the basic domain leucine zipper motifs. Suzuki et al.8 demonstrated that a simple code for DNA recognition can be applied in the case of transcription factors. According to Suzuki et al.,8 the major part of the DNA recognition code consists of two types of rules; chemical and stereochemical. The chemical rules are based on the intrinsic chemical ability of a given residue and a base to form chemical contacts, either a hydrogen bond or hydrophobic interaction in the major groove. The chemical rules making up the DNA recognition code are given in Table 12.1. In addition, the main target for hydrophobic interaction in the major groove of DNA is the single methyl group of the T base, being recognized by hydrophobic residues such as Ala, Val, Ile, Leu, Met, Phe, Tyr, Trp and Thr. Stereochemical rules are summarized in a stereochemical chart which is deduced from crystal and NMR structures of DNA–protein complexes.8 This stereochemical chart indicates which

Table 12.1 Single amino acid–single base contacts (chemical rules). The ‘specific’ residue partners are shown in bold, while nonspecific partners are in plain text. Chemical merit points, semi-arbitrary numbers associated with particular contacts, are used to quantify the energy and specificity of a pairing between an amino acid residue and a nucleotide base. For example, the interaction of Arg (to G), which is particularly favourable and specific, with the residue receives 15 merit points, while the interaction between Ser (to any base), which is less specific, is given 10 points. (M. Suzuki, N. Yagi, DNA-recognition code of transcription factors in the helix-turn-helix,probe helix,hormone receptor, and zinc finger families. Proc. Natl. Acad. Sci, 91, 12357–12361. Copyright 1994, National Academy of Sciences, USA.) small

medium

large

aromatic

A

Cys, Ser, Thr

10

Asn Asp His

15 9 8

Gln Glu Arg, Lys Met

15 9 3 5

Tyr Trp

5 5

T

Ala Cys,Ser, Thr

10

Val, Ile Asn His

12 10 8

Leu, Met Gln Arg, Lys

12 10 5

Tyr, Phe Trp

G

Cys, Ser, Thr

10

His Asn

12 10

Arg, Lys Gln

15 10

Tyr

5

C

Val Cys, Ser, Thr

8 10

Asp Asn His Ile

12 10 8 8

Glu Gln Leu, Met

12 10 8

Tyr, Phe Trp

8 8

12 12

Transition Metal Complex–Peptide Conjugates

349

positions of the protein specifically contact bases and shows what residue sizes are compatible with these positions, resulting in a unique pattern shared by all members of each family of DNA-binding proteins.9 DNA recognition codes were also proposed for other families of DNA-binding proteins such as homeodomains,10 the EGR (early growth response factor) family of zinc-finger transcription factors11 and zinc-finger proteins12 However, it was realized that there is no clear one-to-one correspondence between amino acids and bases.13 Consequently, the rules cannot be generalized for all DNA-binding proteins. Furthermore, a rules-based approach is likely to fail in cases where the same pair interacts using a variety of geometries or a protein recognizes specific DNA sequences indirectly, i.e. via water-mediated contacts, via specific sequence-dependent conformational features and/or via binding-induced distortion of DNA.13 In many cases, peptides have been combined with a metal centre in order to increase their thermodynamic stability and DNA-binding affinity. In this context, research has followed mainly two directions. The first approach involves coordination of a metal ion to a peptide ligand forming metal–peptide complexes. The metal ion may play a structural role imparting structure to an unstructured peptide and/or conferring nucleolytic activity. Many excellent reviews are available with information about the DNA interactions of several types of metal–peptide complexes, such as metal complexes of small peptides,14 lanthanide-based metallopeptides,15 zincfinger peptides,16 chemical nuclease–protein conjugates.17 On the other hand, the second approach focuses on appending the peptide either to an aromatic ring or the backbone of another ligand participating in the coordination of a metal ion, thus forming metal complex–peptide chimeras or conjugates. Parent metal complexes are molecules with established DNA-binding properties. Tethering the peptide to such metal complexes is in most cases anticipated to result in sequence-specificity upon DNA-binding. Transition metal complex–peptide conjugates, discussed herein, offer a new potential in DNA binding and cleavage. They combine the DNA-binding properties of metal complexes accomplished via electrostatic and/or van der Waals interactions, or covalent binding with the ability to form specific contacts with the DNA bases provided by the peptide backbone and side chain donor groups. Additionally, an appropriately designed metal complex may confer its properties to the chimeric molecule resulting in DNA conformational, chemiluminescent or fluorescent probes and chemical nucleases.

12.2 Transition Metal Complex–Peptide Conjugates It is generally known that proteins often use a significant percentage of their amino acids to form the nonspecific contacts that provide the affinity to DNA, while they use an appreciably smaller number of amino acids to make direct contacts with the base pairs. Based on this concept, when constructing metal complex–peptide conjugates, the metal complex provides nonspecific affinity and less steric bulk and complexity than the full DNA binding protein, while the essential recognition site of the original peptide motif is maintained.

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DNA Recognition and Binding by Peptide–Metal Complex Conjugates

12.2.1

Dipeptides

Jan Reedijk and his collaborators have undertaken the construction of a library of peptide-tethered platinum complexes using automated solid-phase synthesis.18 The general structure of the target library is shown in Figure 12.1 and encompasses a dipeptide tethered to an ethylenediamine moiety (en), which, in turn, serves as a platinum chelating ligand. As constituents of the appended dipeptide, six amino acids were selected: glycine, phenylalanine, lysine, arginine, serine and glutamate. Incorporating these amino acids in both positions of the appended dipeptide resulted in a final library of 36 compounds. Subsequently, each compound was screened for in vitro anticancer activity and apparently no important cytotoxicity was observed compared to cisplatin. In continuation of this work, M. S. Robillard et al.19 used the linear amplification system in order to investigate the DNA damage intensity and sequence specificity of adduct formation by six peptide–platinum complexes, cisplatin (1) and Pt(en)Cl2 (2) in pUC19 plasmid as well as in intact Hela cells. The six peptide–platinum complexes were members of the previously described18 library of peptide–dichloroplatinum complexes and differed only in their terminal pendant amino acid: Gly, Phe, Lys, Arg, Ser and Glu. Interestingly, the appended dipeptides did not alter the DNA sequence specificity (runs of consecutive guanines) and did not increase the reactivity with DNA of the complexes compared to the unfunctionalized parent compounds, cisplatin and Pt(en)Cl2. On the other hand, the nature of the terminal amino acid affected the reactivity of the peptide–platinum complexes towards plasmid DNA, with the glycine-tethered complex and the phenylalanine-tethered complex causing the highest damage, followed by (in decreasing order) lysine-tethered, arginine-tethered, serine-tethered and glutamate-tethered. The fact that the negatively charged Glu displayed negligible activity implied that electrostatic interactions may play a role in the DNA binding. 12.2.2 Tripeptides Gly–His–Lys. The tripeptide glycyl-l-histidyl-l-lysine, GHK, has been isolated as a square-planar complex with copper and iron from human plasma and was found to act synergistically with these metals to alter the growth and metabolism of cultured hepatoma cells and hepatocytes.20 The Cu(II) atom is tetracoordinated with the Cl

Cl

O

Pt H2N H2C

NH CH2

C C H2

AA2 N H

O

H N C O

C AA1

NH2

AA = Amino Acid Figure 12.1 The general structure of the dichloroplatinum(II) peptide complexes

Transition Metal Complex–Peptide Conjugates

H5 H 3C 4 m-bpy Lys O H 2N

His O

H CH N

α

C

C

H βC H

O H CH N

α β

CH2

γ

H C H H δC H H εC H

H3

H5 HN 1

C

Gly

H3'

H CH N

C

α

H

O

H5

2 2'

4'

H5'

H6

H6

bpy

H4 N

H3 H3'

2 2'

N

N Ru

2+

H6' H6'

N

N H6'

H6

H5

N

H4

351

2'

H3'

2

H3

H4' H 5' H5' H 4'

bpy

N3 H2

NH2

Figure 12.2 Structure of the complex D-[Ru(bpy)2(m-bpy-GHK)]2+ with atom numbering

imidazole nitrogen, a peptide nitrogen, the terminal glycyl amino group, and an oxygen atom from a water molecule.21,22 In order to investigate its possible sequence specificity upon DNA binding, N. Hadjiliadis and collaborators23,24 conjugated peptide GHK to the bpy ligand of the parent complex [Ru(bpy)3]2+ (bpy = 2,2′-bipyridine) (Figure 12.2) which is known to bind DNA reversibly25,26 and induce photoactivated cleavage.27 A 1H NMR study of the interaction of the resulting diastereomeric complexes L- and D-[Ru(bpy)2(mbpy-GHK)]2+ (m-bpy = 4-carboxy-4′-methyl-2,2′-bipyridine) with two different DNA sequences, the d(CGCGAATTCGCG)223 and d(CGCGATCGCG)2,24 revealed that minor changes in the DNA sequence can result in different binding characteristics. In the case of the system D-[Ru(bpy)2(m-GHK)]2+–d(CGCGAATTCGCG)2 1:1, molecular models based on the NOE contacts (Figure 12.3) showed that the Disomer bound to the DNA major groove with the two bpy ligands oriented towards the double helix. The L-isomer, on the other hand, approached the DNA minor groove using the m-bpy ligand placing the lysine ε-amino group close to the DNA phosphates (Figure 12.3).23 Furthermore, significant chemical shifts of peptide and cytosine’s exocyclic amino group protons, as well as several intermolecular NOEs demonstrated that the peptide GHK was oriented towards the DNA major groove close to the central AATT sequence forming possibly specific contacts with the DNA bases. However, no significant interaction was observed for the D-[Ru(bpy)2(mGHK)]2+–d(CGCGATCGCG)2 system at ratio 1:1. Interestingly, increase of the metal complex to DNA duplex ratio resulted in binding to the major groove and strand opening of the DNA duplex. At 2:1 ratio, the majority of the intermolecular NOEs took place between the peptide Gly–His–Lys and sugar protons located in the major groove. This observation implied an important role for the peptide upon

352

DNA Recognition and Binding by Peptide–Metal Complex Conjugates

Figure 12.3 Molecular models of the interactions between [d(CGCGAATTCGCG)2] and D[Ru(bpy)2(m-GHK)]Cl2 (A) and L-[Ru(bpy)2(m-GHK)]Cl2 (B) reflecting the average binding of each complex to the oligonucleotide. (A. Myari, N.Hadjiliadis, A.Garoufis, Synthesis and characterization of the diastereomers L- and D-[Ru(bpy)2(m-bpy.Gly-His-L-His-L-Lys)]Cl2. 1H NMR studies on their interactions with the deoxynucleotide duplex d[5′-CGCGAATTCGCG3′], Eur. J. Inorg. Chem., 2004, 7, 1435–1436. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Reproduced with permission.)

interaction of the chimera with DNA, but the absence of significant shifts of the peptide functional groups (or their neighbouring protons) indicated that probably no specific contact between peptide and the DNA bases was formed, rather an electrostatic type of interaction was assumed. Photocleavage studies were also conducted with the plasmid pUC19 and a 158 bp restriction fragment showing that both diastereomers cleave DNA with similar efficiency, attacking mainly the guanines of the sequence probably by generating active oxygen species.24 In an attempt to combine the notable cytotoxicity observed for chloro(polypyridyl) ruthenium complexes28 with the possible DNA recognition specificity provided by the peptide moiety, K. Karidi et al. synthesized the chloro-complex of the tethered peptide [Ru(terpy)(4-CO2H-4′-Mebpy-Gly-l-His-l-LysCONH2)Cl](PF6).29 The interactions of the ruthenium peptide complex with CT-DNA and plasmid DNA were studied with various spectroscopic techniques, showing coordinative binding of the ruthenium complex to DNA, after hydrolysis of the coordinated chloride, and a preference for the bases guanine and cytosine over the bases thymine and adenine. Electrostatic interactions between the complex cation and the polyanionic DNA chain assisted this binding whereas DNA unwinding occurred in all cases with high binding ratio (Ru/base) values (r > 0.3). Moreover, interaction of the peptide with DNA was evidenced by 31P NMR spectroscopy for the one of the two positional isomers of the ruthenium complex.

Transition Metal Complex–Peptide Conjugates

353

Numerous ruthenium nitrosyl complexes containing polypyridyl ligands have been synthesized, with possible uses as regulators of blood pressure30,31 or as antitumour agents releasing NO within tumour cells.32,33 Ruthenium nitrosyls may photorelease the coordinated NO in aqueous media.34,35 The complex [RuII(terpy)(4COGHK-4′-Mebpy)(NO)] was found to be stable only in nonaqueous solvents, and in contact with the water it is transformed into the nitro derivative.36 Photoactivation with visible light of the nitrosyl complex in dry MeCN resulted in the release of NO, while there was partial release of NO2− from the nitro complex. A DNA-binding study was performed in order to investigate the possible role of the peptide GHK using the nitro complex, since it was the only one stable in aqueous media. Interaction of the nitro complex with short fragments (70–300 bp) of calf thymus DNA induced slight shortening of the apparent polynucleotide length, while the peptide GHK was found to interact with the DNA helix in a synergistic way with the whole complex.36 Gly–Gl–Ser. Recently, M. J. Hannon et al.37 explored the possibility of attaching peptides to metallosupramolecular cylinders in order to create sequence-specific DNA binding agents targeted to the less-exploited DNA major groove. The parent cylinders were tetracationic triple stranded (Fe(II)) and dicationic double-stranded (Cu(I) or Ag(I)), based on two bis-pyridylimine ligands with the peptide functionality placed at position 5 of the pyridine ring (termed Lp1 and Lp2, Figure 12.4). The peptide selected for conjugation to the cylinder was the tripeptide Gly-Gly-Ser-CoNH2. The two glycine residues were selected in order to provide the minimal steric bulk close to the cylinder, whereas the serine residue at the Cterminal was chosen based on its known frequent occurrence in protein motif–DNA base contacts. The parent tetracationic triple-stranded iron(II) cylinder binds with CT-DNA in the major groove, inducing dramatic intramolecular DNA coiling38 in the heart of three-way and other Y-shaped junctions.39a,b The parent Cu(I) dicationic cylinder also binds CT-DNA and in the presence of hydrogen peroxide induces H2 C H C H2NOC-Ser-Gly-Gly

H N

C

N

N

N

N

Lp1 CH3 H C

C O

N

H3C

5

C

H N

Gly-Gly-Ser-CONH2

CH2 N H2 C

Lp2

6

4

H3 C

N CH2

3

O

H2 C

H2 C

H2NOC-Ser-Gly-Gly

2 1

O

H N

H C

H C

2 1

CH3

N

3

6

4 5

C

H N

Gly-Gly-Ser-CONH2

O

Figure 12.4 The bis-pyridylimine based, bis-peptide end functionalised ligands, Lp1 and Lp2 with atom numbering at the peptide-substituted pyridine ring

354

DNA Recognition and Binding by Peptide–Metal Complex Conjugates

DNA cleavage.39c DNA-binding experiments of the peptide-functionalized cylinders indicated that the Fe(II) cylinder retained its ability to coil DNA, albeit to less extent compared to the unsubstituted analogue, whereas the Cu(I) cylinder cleaved DNA, with a tendency for double-stranded cleavage.37 Apparently, this is a promising strategy for the construction of sequence-specific major-groove-binding molecules by attaching peptides able to confer sequence-specificity upon DNA binding. 12.2.3

Bioactive Peptides

J. Brunner and J.K. Barton have addressed the problem of insufficient cellular uptake of drugs by attaching a cell-penetrating peptide onto the bpy (2,2′bipyridine) ligand of the rhodium intercalator [Rh(chrysi)(phen)(bpy)]3+ (chrysi = chrysenequinone diimine, phen = 1,10-phenanthroline).40 d-octaarginine, a well-known cell-penetrating peptide for delivery to the nucleus,41 was appended to the rhodium complex. Rhodium intercalators bearing the sterically demanding ligand chrysi bind preferentially to single-base mismatched sites and, upon photoactivation, the complexes furthermore promote strand breakage neighbouring the mismatched site.42 The presence of the oligoarginines was found to increase the nonspecific binding affinity (at least two orders of magnitude) of the complexes for both matched and mismatched DNA, while photocleavage remained selective for the mismatched site. Confocal microscopy experiments confirmed the nuclear localization of the metal–peptide conjugates demonstrating that the delivery properties of the appended cell-penetrating peptide are not affected by the presence of the bulky octahedral metal complex. 12.2.4

De novo Designed Peptides

The variety of the different amino acid side chain functionalities offers a great tool in designing peptides of diverse composition in order to elucidate peptide–DNA interactions. K.D. Copeland et al. used metallointercalator–peptide conjugates as a model system for studying DNA–protein crosslinking.43 The ruthenium intercalator, [Ru-(phen)(bpy′)(dppz)]2+ (bpy′ = 4-(butyric acid)-4′-methyl-2,2′-bipyridine), binds with high affinity to DNA and initiates crosslinking through a flash-quench reaction. Pentamer peptides were designed in order to be long enough to allow interactions between the peptide and DNA, even when the dppz ligand is intercalated into the base stack, and short enough to remain unstructured. An Ala was added at the Ntermini intended as a flexible spacer, and a Gly at the C-termini simplified synthesis and was intended to adsorb end effects. Finally, the three central residues were varied. Two types of peptides were considered: peptides with positively charged residues (Lys or Arg) at positions 2 and 4 and peptides with aliphatic residues (Ala and Gly) at positions 2 and 4. The outcome of this study was that the peptide composition does impact binding and crosslinking. The conjugates bearing lysine residues generally showed greater affinity for DNA. Furthermore, the Lys–X–Lys or Arg–X–Arg peptides gave the highest levels of crosslinking. This difference in crosslinking activity may be attributed to increased interaction between the DNA

Transition Metal Complex–Peptide Conjugates

355

and the positively charged peptide or to the reactivity of the Lys residues. Variation of the central residue did produce some change in binding affinity. The binding constants increased in the order Ru-KWK < Ru-KAK < Ru-KYK. Even though the Trp residue was expected to contribute more in binding affinity through intercalation, probably it was not ideally positioned since tethered peptides are constrained. The differences observed in crosslinking were quite small as the central position in the peptide was varied. 12.2.5

Protein Fragments

In some cases, intact DNA-binding protein fragments with individual recognition properties were chosen for conjugation to the transition metal complexes. The aim was to construct a sequence-specific chimera in which the peptide recognizes its cognate DNA sequence. M.Y. Ogawa and his coworkers44 prepared a chimeric metallo-bZIP protein (Figure 12.5), (GBR-CC)Ru, in which [Ru(bpy)2(phen-IA)]2+ (phen-IA = N-

Figure 12.5 Computer generated model of the synthetic metalloprotein of the designed photoactive DNA binding peptide developed from the crystal structure of cFos-cJun bound to DNA. (Reprinted from Inorg. Chim. Acta, 300–302, Lasey R.C, Banerji S.S., Ogawa M.Y., Synthesis and characterization of a sequence-specific DNA-binding protein that contains ruthenium polypyridyl centres, 822–828. Copyright 2000, with permission from Elsevier.)

356

DNA Recognition and Binding by Peptide–Metal Complex Conjugates

iodoacetyl-5-amino-1,10-phenanthroline) was attached to the coiled-coil (CC) dimerization region based on an artificial sequence developed by Hodges and coworkers (Figure 12.5).45 The DNA-binding domain of the protein was taken from the basic region of the GCN-4 transcription factor (GBR). The photoactive metal complex was attached to Cys29, the most heavily solvent-exposed position of the coiled-coil closest to the DNA-binding domain, and, as indicated by its emission spectrum, the ruthenium centre remained exposed to solvent when coupled to the bZIP peptide. Electrophoretic mobility shift assays showed that the chimeric metalloprotein retained the essential DNA recognition properties of the native GCN-4 transcriptional activator. Peptide–DNA complexes were formed with the AP1 and CRE sequences, known recognition sites of GCN-4, but not the divergent Sp1 sequence. Peptide titration studies indicated that the affinities of (GBR-CC)Ru for the AP1 and CRE sites were comparable. (GBR-CC)Ru did not produce observable photoinduced DNA damage, probably due to the separation distance between the ruthenium sites and the DNA bases.44 The DNA recognition properties of the a3-helix of the DNA-binding phage P22 repressor were investigated by attachment onto the metallointercalating [Rh(phi)2(phen′)]3+ (phi = 9,l0-phenanthrenequinone diimine; phen′ = 5(amidoglutaryl)-1,10-phenanthroline).46 Remarkably, a single glutamate at position 10 was found to be essential in directing DNA site-recognition to the sequence 5′-CCA-3′. It is noteworthy that conservative modifications of this glutamate, including aspartate substitution or derivatization to the glutamate methyl ester, abolished the 5′-CCA-3′ recognition. On the other hand, circular dichroism indicated significant a-helical content in the peptide, which depended upon the presence of the glutamate. The experimental evidence indicated that the glutamate may play a dual role, folding the metal–peptide complexes into a unique conformation and also interacting directly with DNA. However, the metal–peptide complexes did not preferentially target the putative operator sequence for protein P22R. This fact may arise because in the native protein, intraprotein interactions determine the orientation of the a3 recognition helix, whereas in the metal–peptide complexes, the peptide conformation may be directed instead by the metal centre and the solvent. Furthermore, moving the glutamate at position 10 in the sequence of the appended peptide to position 6 changed the sequence preference of the metallointercalator–peptide conjugate to 5′-ACA-3′, whereas replacing Glu6 with Arg resulted in a change in the apparent consensus sequence from 5′-ACA-3′ to 5′-(T/G)CA-3′.47 These results clearly illustrate that with a single amino-acid modification, the recognition sequence and the photocleavage characteristics of the metallointercalator–peptide conjugate can be perturbed. A similar work by N.Y. Sardesai and J.K. Barton focused on the DNA recognition properties of metal–peptide complexes containing peptides derived from the sequence of the a3 recognition helix of the phage 434 repressor protein (434R) appended to the metallointercalator [Rh(phi)2(phen′)]3+.48 A series of peptide sequences was designed, based upon 434R and criteria for maximizing peptide ahelicity. The three Gln residues were left unchanged because they make base-specific contacts with the 434R consensus sequence 5′-ACAA-3′. Therefore, a family of peptides aA, aB (aA–Q7R), aC (aA–Q3R), and aD (aA–Q3A) was formed, in which peptides were different from one another by a single amino acid at one of the three

Transition Metal Complex–Peptide Conjugates

357

Gln residues implicated in DNA recognition. The 5′-ACAA-3′ sites were cleaved strongly by all metal–peptide complexes. However, in contrast to 434R, they showed a slight 5′-ACGA-3′ > 5′-ACAA-3′ preference. Single-site mutants at Q7 or Q3 did not affect the relative specificity of the metal–peptide complexes, but changed the affinity. Two issues were considered to contribute in the reduction in specificity. The first issue involved the intrinsic site selectivity of the metallointercalator resulting in preference also for the sites 5′-ACA-3′ or 5′-ATG-3′. The second issue addressed the conformational flexibility of appended peptides resulting in decrease of the thermodynamic advantage to be gained from the ensemble of contacts available in the DNA-binding protein or even promote nonspecific interactions with the phosphate backbone, so that the major contribution to DNA site specificity would be derived instead from the metal complex.48 Another interesting study involved a peptide (7p) derived from the recognition loop of the restriction endonuclease MunI from Mycoplasma unidentified, which was appended to the groove-binding complex [Ru(bpy)2(m-bpy)]2+ (m-bpy = 4methyl-4′-carboxylate-2,2′-bipyridine).49 MunI recognizes the palindromic hexanucleotide sequence C/AATTG and cleaves as indicated by the slash. All amino acid residues involved in sequence-specific interactions lie within a single short region (Arg115-Gly116-Asn117-Ala118-His119-Glu120-Arg121 = 7p).50 A 1H NMR study of the interactions of the diastereomeric complexes L- and D-[Ru(bpy)2(m-bpy-7p)]2+ with the oligonucleotide sequences d(5′-CGCGATCGCG-3′)2 and d(5′-GCGCTTAAGCGC-3′)2 indicated binding of both isomers to the ends of both sequences. Furthermore, the L-isomer exhibited additional NOE contacts to the major groove protons of the GCT/CGA sequence of the decanucleotide and the central TT/AA part of the dodecanucleotide. No sequence-specific contacts were indicated by the experimental results, albeit intermolecular NOEs showed that the peptide was oriented towards the DNA major groove. Thus, the appended peptide did not reproduce the recognition characteristics of the endonuclease that it was derived from. This is another case in which the conformational flexibility of the peptide moiety did not allow formation of sequence-specific contacts, whereas in the case of d(5′GCGCTTAAGCGC-3′)2 the resultant conformers were bound to DNA via many different orientations.49

12.2.6

Summary

Tethering of a peptide to a transition metal complex, especially to a metallointercalator, increases the overall DNA-binding affinity. Many studies have shown that minimalist peptide domains are insufficient for tight binding and dimerization has been utilized in order to enhance affinity.7 N.Y. Sardesai et al. have demonstrated that a 100-fold excess of free peptide was incapable of competing with the corresponding rhodium metallointercalator-peptide conjugate in binding to the DNA target.46 On the other hand, the peptide contributes substantially to the DNAbinding affinity, resulting in tighter binding compared to the metal complex.48 Simple coordination metal complexes often lack hydrogen-bonding donors or acceptors, thus undergoing their site-specific reactions solely on the basis of shapes and symmetries.51 Thus, in most cases, the appended peptide contributed

358

DNA Recognition and Binding by Peptide–Metal Complex Conjugates

significantly in the sequence specificity of the conjugates, being able to discriminate between different DNA sequences. Furthermore, the sequence selectivity was in many cases perturbed by a single amino acid modification,46,47 demonstrating the versatility of peptides as systems utilized for DNA recognition. However, in some cases there is a possibility that the intrinsic preference of the metal complex for a specific DNA sequence may interfere with sequence specificity conferred by the peptide moiety. Hence, the resulting sequence specificity may differ from that expected in cases where the peptide sequence is derived from a DNA-binding protein.48 Although, small peptides are composed of readily available building blocks and it is facile to tune the peptide composition to achieve the desired properties and variations, they lack the structural constraints and control that may be needed for predictive design. Indeed, their conformational flexibility accounts for the failure of recognizing the same sequence as the native protein they are derived from.48,49

12.3

Metallointercalator–Metallopeptide Conjugates

Design of tethered peptides has also been extended to mimicking the properties of certain proteins, such as DNA hydrolases, zinc-binding proteins and other important cell proteins which interact with DNA. Barton and coworkers tethered a peptide designed de novo, based on the active sites of metal-containing hydrolases to the metallointercalator [Rh(phi)2(bpy′)]3+.52 Two histidines were placed in positions 7 and 11 along a 16-residue peptide (P1) to create a zinc coordination site on one face of an a-helix. A glutamate was included at position 4, and glutamate–lysine salt bridges, as well as space-filling alanines were added to increase a-helicity (Figure 12.6). Both helical content and cleavage activity of plasmid DNA reached a maximum at stoichiometric amounts of Zn2+ using micromolar concentrations of the chimeric complex, owing to the strong binding affinity of the intercalating moiety. Furthermore, direct evidence was obtained for a hydrolytic cleavage reaction.52 Interestingly, the same peptide, when conjugated to ethidium, in the presence of Zn(II), forms a closed chelate structure which prevents binding to DNA and concomitant cleavage.53 The importance of several key residues of P1 was further assessed by determining the DNA cleavage efficiency of mutations in plasmid cleavage assays.54 Not surprisingly, the two histidine residues at positions 7 and 11 were found to be important for zinc coordination and cleavage activity. On the other hand, the cleavage rate constant was not affected upon mutation of the glutamic acid at position 4.54 Also, the selection of the metallointercalator seemed to be critical for the hydrolytic activity, since the attachment of P1 to [Rh(phen)(bpy)(chrysi)]3+, a metallointercalator which binds to DNA mismatches, did not produce Zn2+-promoted hydrolysis.54 In addition, zinc(II)-promoted cleavage of supercoiled plasmid has also been demonstrated for another metallointercalator–metallopeptide conjugate bearing a short b-hairpin derived from the BamHI restriction endonuclease, incorporating the three important catalytic residues (Asp94, Glu111 and Glu113) responsible for the hydro-

A Critical Survey and Future Perspectives phi

359

3+

NH HN

H N phi

Rh N H

N N

bpy H N Asp Pro Asp Glu Leu Glu His Ala Ala Lys His Glu Ala Ala Ala Lys CONH2 O HN

NH N

N Zn 2+

Figure 12.6 The Rh(III) complex-peptide conjugate, Rh(phi)2bpy′-Peptide (Rh-P1) with the Zn(II) coordination site being illustrated

lytic activity of the native enzyme.54 It must be emphasized that both appended peptides did not contribute to the recognition properties of the sequence-neutral metallointercalator [Rh(phi)2(bpy′)]3+ but efficiently delivered the metal ion to the sugar-phosphate backbone.

12.4 A Critical Survey and Future Perspectives Research on the development of artificial nucleases aims at high cleavage efficiency, DNA affinity and sequence selectivity. Attempting to compare different transition metal complex–peptide conjugates regarding their DNA cleavage efficiency becomes a rather difficult task due to the large variety of reaction conditions used. Therefore, comparisons can be made tentatively under the concept that in most cases, reaction conditions are optimized in order to give the best results. Furthermore, there is no available data sufficient for comparisons of kinetic parameters. Experimental data derived from the studies already described are summarized in Table 12.2. First of all, DNA binding by intercalation increases DNA binding affinity by at least two orders of magnitude compared to DNA groove binding. For example, for the series [Ru(bpy)3]2+, [Ru(bpy)2(phen)]2+, [Ru(bpy)2(DIP)]2+ and [Ru(bpy)2(phi)]2+, Kb values of 0.7 × l03, 0.7 × l03, 1.7 × l03, and 1.6 × l05 M−1, have been determined respectively.55 Given that increased DNA binding affinity results, in most cases, in high DNA cleavage efficiency, it is clearly seen from Table 12.2 (cleavage efficiency is compared as % formation of nicked and linear DNA from plasmid DNA

hydrolytic oxidative hydrolytic hydrolytic – hydrolytic hydrolytic hydrolytic oxidative oxidative oxidative oxidative

[Rh(phi)2(bpy′)]-P1 + Zn(II) 1:1 [Rh(phi)2(bpy′)]-P1 + Cu(II) 1:1 [Rh(phi)2(bpy′)]-P1 + Cd(II) 1:1 [Rh(phi)2(bpy′)]-P1 + Fe(II) 1:1 [Rh(phi)2(bpy′)]-Bamb [Rh(phi)2(bpy′)]-Bam + 100 eq Zn(II) [Rh(phi)2(bpy′)]-Bam + 100 eq Cd(II) [Rh(phi)2(bpy′)]-Bam + 100 eq Mg(II) [Rh(phi)2(bpy′)]-Bam + 1 eq Cu(II) [Rh(phi)2(bpy′)]-Bam + 1 eq Fe(II) [Rh(phi)2(phen′)]-aAc [Rh(phi)2(phen′)]-PAd 44 45 50 20 38 54 57 41 62 35

(18) (18) (18) (18) (26) (26) (26) (26) (26) (26) – –

65 (10) 40 (18)

% nicked plasmid DNAe

† Cleavage sites are underlined. * Samples have been irradiated. nd Sequence selectivity has not been determined. a P1 is a de novo designed peptide incorporating histidine residues for Zn2+ coordination. b Bam is a peptide derived from the active center of BamHI. c aA is a peptide derived from the recognition helix of phage 434 repressor protein. d PA is a peptide derived from the a3 helix of the phage P22 repressor. e The control experiment (plasmid only) is given in parenthesis.

oxidative –

[Ru(bpy)2(m-GHK)]2+ [Rh(phi)2(bpy′)]-P1a

Mechanism of cleavage

38 (0) – – – – 2 (0) 4 (0) – 3 (0) 2 (0) – –

– 4 (0)

% linear plasmid DNAe

5′-NACAA-3′* 5′-CCA-3′*

nd

nd

nd

nd

nd

nd

nd

nd

Mainly G* Modest selectivity for 5′-Pu-Py-Pu-3′* 5′-Py-Py-Pu-Pu-Pu-3′ 5′-Py-Py-Pu-Pu-Pu-3′

Sequence selectivity†

Table 12.2 Nuclease activity of transition metal complexes-peptide conjugates on super-coiled plasmid

54 54 54 54 54 54 54 54 54 54 48 46

24 54

Reference

360 DNA Recognition and Binding by Peptide–Metal Complex Conjugates

A Critical Survey and Future Perspectives

361

degradation) that the [Ru(phi)2(bpy′)]-P1 + Zn(II) 1:1 [54] (where P1 is a de novo designed a-helical peptide containing histidine residues for Zn2+ coordination) is the most efficient DNA cleaving metal complex with totally 82% formation of nicked plasmid and linear DNA (64% increase of modified plasmid DNA compared to control sample). On the other hand, high sequence selectivity is obtained in the case of appended peptides derived from DNA-binding proteins, especially when these peptides are either already structured (e.g. a3 helix of the phage P22 repressor46) or initially designed to be a-helical, with a-helicity further increased upon metal coordination (e.g. coordination of Zn(II) to appropriately designed peptides54). However, it is important to keep in mind that sequence selectivity and cleavage efficiency of intercalating metal complexes tethered to peptides can be further improved and optimized by the selection of the appropriate ancillary ligands, intercalating ligands, linker and appended peptides. Metal–peptide complexes show higher cleavage ability of DNA compared to metal complex–peptide conjugates. As seen from Table 12.2, it is obvious that 100% cleavage of plasmid DNA is not reported for any of the metal complex–peptide conjugates (82% is the highest percentage of cleavage observed in the case of [Rh(phi)2(bpy′)]-P1+Zn(II) 1:1). On the other hand, 100% conversion of supercoiled plasmid DNA (form I) to nicked circular DNA (form II) and linear DNA (form III) is reported for most metal-coordinated peptides.56–58 Furthermore, Ni(II) metallopeptides of the type Xaa–Xaa–His show remarkably high DNA-binding constants,56 comparable with those of intercalating metal complexes tethered to peptides (107–108 M−1).46,47 Since metal complex–peptide conjugates and Ni(II) metallopeptides bind to DNA with similar affinities, then the reason for the higher cleavage efficiency observed for Ni(II) metallopeptides probably lies in the different mechanisms of DNA cleavage. A possible explanation is the shape selectivity upon DNA binding of Ni(II) metallopeptides, reminiscent of the known minor-groove binding drugs, that enables them to fit into the DNA minor groove bringing the metal centre in close proximity to the deoxyribose moiety (cleavage takes place commonly by C4′-H abstraction). Even higher cleavage efficiency can be accomplished when metallopeptides are conjugated to an intercalating ligand, such as in the case of (bis-Ni(II)·GGH-NDI), in which two Ni(II)·GGH units are conjugated to napthalene diimide (NDI).59 The resultant nuclease is the most efficient reported so far among metallopeptides (100 times more efficient than Ni(II)·GGH).59 In addition, sequence selectivity and cleavage efficiency of metal–peptide complexes can conveniently be modulated either by changing the position of amino-acids bearing the appropriate side chain functionalities within the peptide sequence, thus discriminating between different mixed A/T sequences or changing amino acid chirality resulting in totally different sequence recognition (e.g. 5′-CCT-3′ in the case of Ni(II)·Gly-D-Asn-His56). Another important issue concerning the design of applicable artificial nucleases is the mechanistic pathway followed. Formation of diffusible active oxygen species results in base-directed reactivity, with guanines being the most vulnerable residues toward oxidation. Consequently, any sequence selectivity observed upon DNA binding cannot be reproduced after initiation of cleavage. This is obvious in the case of peptides conjugated to the parent complex [Ru(bpy)3]2+.24 Hence, formation of

362

DNA Recognition and Binding by Peptide–Metal Complex Conjugates

diffusible radicals upon DNA cleavage is undesirable when designing an artificial nuclease bearing high sequence selectivity. Often, a hydrolytic rather than an oxidative cleavage mechanism is desirable for several reasons. Production of diffusible free radicals by oxidative cleavage of DNA results in strand ends that cannot be enzymatically religated in molecular biology applications. For clinical applications, oxidative cleavage can cause indiscriminate peripheral damage to the cell while radical diffusion may affect significantly the specificity of cleavage that can be achieved.15 Rh(III) metallointercalator– Zn(II) metallopeptide conjugates are the only chimeric molecules utilizing a hydrolytic mechanism of cleavage (Table 12.2). The pseudo first-order rate constant of [Rh(phi)2(bpy′)-P1+Zn(II) (1 × 10−5 s−1)54 determined for plasmid DNA degradation is of the same magnitude to the bimetallic Zn(II)2-heptapeptide developed by P. Scrimin et al.58 and to lanthanide metallopeptides.60 Therefore, conjugation of a Rh(III) metallointercalator to a hydrolytically active Zn(II) metallopeptide appears to be an effective system for the hydrolysis of DNA, displaying moderate sequence selectivity.54 The versatility of such systems could probably meet the requirements for obtaining the desired hydrolytic activity in the near future.

12.5

Conclusion

By careful design of transition metal complex–peptide conjugates, specific recognition of a DNA site may be achieved, which in some cases can be perturbed by a single amino acid modification. However, there are some limitations for the recognition of a DNA-binding protein cognate sequence by a peptide model of the native protein. In this case, issues of peptide flexibility need to be addressed. Indeed, highly structured appended peptides provide a higher degree of specificity for DNA recognition. DNA specific recognition is often accompanied by significant nucleolytic activity. Transition metal complex–peptide conjugates are not as efficient in DNA cleavage as metal-coordinated peptides. The reason probably lies in the different mechanism employed in each case. However, appropriately designed appended peptides may result in significant DNA hydrolytic activity by delivering Zn(II) ions to the DNA phosphate backbone. Importantly, metal complex–peptide conjugates are able to interact with the DNA major groove, commonly exploited upon DNA recognition by DNA-binding proteins. The versatility of these systems has become evident, thus further enhancement of their DNA recognition and cleavage properties may be achieved by appropriate selection of the ancillary ligands, the peptide composition and the metal ion.

Abbreviations AP1 bpy

Activator protein 2,2′-Bipyridine

References

bZip chrysi CRE CT-DNA DIP dppz en GGH GHK KAK KWK KYK m-bpy NDI NOE Phen-IA terpy

363

Basic leucine zipper Chrysenequinone diimine Cyclization recombination Calf thymus deoxyribose nucleic acid 4,7-Diphenyl-1,10-phenanthroline Dipyridophenazine Ethylene diamine Glycyl–glycyl–histidine Glycyl–histidyl–lysine Lysyl–alanyl–lysine Lysyl–tryptophanyl–lysine Lysyl–tyrosyl–lysine 4-Carboxy-4′-methyl-2,2′-bipyridine Napthalene diimide Nuclear Overhauser Effect N-iodoacetyl-5-amino-1,10-phenanthroline 2,2′:6′,2″-Terpyridine

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8.

9. 10. 11. 12. 13. 14. 15. 16.

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21.

DNA Recognition and Binding by Peptide–Metal Complex Conjugates Chem. Soc., 1994, 116, 11137–11138. (c) Morii, T.; Yamane, J.; Aizawa, Y.; Makino, K.; Sugiura, Y.; Cooperative oligomerization enhances sequence-selective DNA binding by a short peptide; J. Am. Chem. Soc., 1996, 118, 10011–10017. (a) Talanian, R.V.; McKnight, C.J.; Kim, P.S.; Sequence-specific DNA binding by a short peptide dimer; Science, 1990, 249, 769–771. (b) Morii, T.; Simomura, M.; Morimoto, S.; Saito, J.; Sequence-specific DNA binding by a geometrically constrained peptide dimer; J. Am. Chem. Soc., 1993, 115, 1150–1151. (a) Benos, P.V.; Lapedes, A.S.; Stormo, G.D.; Is there a code for protein–DNA recognition? Probab (ilistical)ly …; BioEssays, 2002, 24, 466–475. (b) Suzuki, M.; Brenner, S.E.; Gerstein, M.; Yagi, N.; DNA recognition code of transcription factors; Prot. Eng., 1995, 8, 319–328. Suzuki, M.; Yagi, N.; DNA recognition code of transcription factors in the helix-turnhelix, probe helix, hormone receptor and zinc finger families; Proc. Natl Acad. Sci. USA, 1994, 91, 12357–12361. Damante, G; Pellizzari, L.; Esposito, G.; Fogolari, F.; Viglino, P.; Fabbro, D.; Tell, G.; Formisano, S.; Lauro, R.D.; A molecular code dictates sequence-specific DNA recognition by homeodomains; EMBO J., 1996, 15, 4992–5000. Benos, P.V.; Lapedes, A.S.; Stormo, G.D.; Probabilistic code for DNA recognition by proteins of the EGR family; J. Mol. Biol., 2002, 323, 701–727. Sera, T.; Uranga, C.; Rational design of artificial zinc-finger proteins using a nondegenerate recognition code table; Biochemistry, 2002, 41, 7074–7081. Sarai, A.; Kono, H.; Protein–DNA recognition patterns and predictions; Annu. Rev. Biophys. Biomol. Struct., 2005, 34, 379–398. (a) Long, E.C.; Ni(II)·Xaa-Xaa-His metallopeptide-DNA/RNA interactions; Acc. Chem. Res., 1999, 32, 827–836. (b) Mancin, F.; Scrimin, P.; Tecilla, P.; Tonellato, U.; Artificial metallonucleases; J. Chem. Soc. Chem. Commun., 2005, 2540–2548. (a) Franklin, S.J.; Lanthanide-mediated DNA hydrolysis; Curr. Opin. Chem. Biol., 2001, 5, 201–208. (b) Franklin, S.J.; Welch, J.T.; The helix-turn-helix as a scaffold for chimeric nuclease design; Comm. Inorg. Chem., 2005, 26, 127–164. (a) Beerli, R.R.; Barbas, III, C.F.; Engineering polydactyl zinc-finger transcription factors; Nat. Biotechnol., 2002, 20, 135–141 and refs therein. (b) Pabo, C.O.; Peisech, E.; Grant, R.A.; Design and selection of novel Cys2His2 zinc finger proteins; Ann. Rev. Biochem., 2001, 70, 313–340. (c) Jantz, D.; Amann, B.T.; Gatto, Jr, G.J.; Berg, J.M.; The design of functional DNA-binding proteins based on zinc finger domains; Chem. Rev., 2004, 104, 789–800. (d) Dhanasekaran, M.; Negi, S.; Sugiura, Y.; Designer zinc finger proteins: tools for creating artificial DNA-binding functional proteins; Acc. Chem. Res., 2006, 39, 45–52, (e) Durai, S.; Mani, M.; Kandarelou, K.; Wu, J.; Porteus, M.H.; Chandrasegaran, S.; Zinc finger nucleases: custom-designed molecular scissors for genome engineering of plant and mammalian cells; Nucl. Acids Res., 2005, 33, 5978–5990. Sigman, D.S.; Bruice, T.W.; Mazumder, A.; Sutton, C.; Targeted chemical nucleases; Acc. Chem. Res., 1993, 26, 98–104. Robillard, M.S.; Bacac, M.; Elst, H.; Flamigni, A.; van der Marel, G.A.; Van Boom, J.H.; Reedijk, J.; Automated parallel solid-phase synthesis and anticancer screening of a library of peptide-tethered platinum(II) complexes; J. Comb. Chem., 2003, 5, 821–825. Robillard, M.S.; Davies, N.P.; van der Marel, G.A.; Van Boom, J.H.; Reedijk, J.; Murray, V.; The interaction of peptide-tethered platinum(II) complexes with DNA; J. Inorg. Biochem., 2003, 96, 331–338. (a) Pickart, L.; Thaler, M.M.; Tripeptide in human serum which prolongs survival of normal liver cells and stimulates growth in neoplastic liver; Nature New Biol., 1973, 243, 85–87. (b) Schlesinger, D.; Pickart, L.; Thaler, M.M.; Growth modulating serum tripeptide is glycyl-histidyl-lysine; Experientia, 1977, 33, 324–325. (c) Pickart, L.; Thaler, M.M.; Millard, M.M.; Effect of transition metals on recovery from plasma of the growthmodulating tripeptide glycylhistidyllysine; J. Chromatogr., 1979, 175, 65–73. Freedman, J.H.; Pickart, L.; Weinstein, B.; Mims, W.B.; Peisach, J.; Structure of the glycylL-histidyl-L-lysine-copper(II) complex in solution; Biochemistry, 1982, 21, 4540–4544.

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22. Laussac, J.P.; Haran, R.; Sarkar, B.; N.m.r. and e.p.r. investigation of the interaction of copper(II) and glycyl-L-histidyl-L-lysine, a growth-modulating tripeptide from plasma; Biochem. J., 1983, 209, 533–539. 23. Myari, A.; Hadjiliadis, N.; Garoufis, A.; Synthesis and characterization of the diastereomers L- and D-[Ru(bpy)2(m-bpy-Gly-L-His-L-Lys)]Cl2 – 1H NMR studies on their interactions with the deoxynucleotide duplex d[(5′-CGCGAATTCGCG-3′)2]; Eur. J. Inorg. Chem., 2004, 1427–1439. 24. Myari, A.; Hadjiliadis, N.; Garoufis, A.; Malina, J.; Brabec, V.; NMR analysis of duplex d(CGCGATCGCG)2 modified by L- and D-[Ru(bpy)2(m-GHK)]Cl2 and DNA photocleavage study; J. Biol. Inorg. Chem., 2006, 12, 279–292. 25. Yamagishi, A.; Evidence for stereospecific binding of tris(1,10-phenanthroline)ruthenium(II) to DNA is provided by electronic dichroism; J. Chem. Soc. Chem. Commun., 1983, 572–573. 26. Yamagishi, A.; Electric dichroism evidence for stereospecific binding of optically active tris chelated complexes to DNA; J. Phys. Chem., 1984, 88, 5709–5713. 27. Sigman, D.S.; Mazumder, A.; Perrin, D.M.; Chemical nucleases; Chem. Rev., 1993, 93, 2295–2316. 28. Novakova, O.; Kasparkova, J.; Vrana, O.; Vanvliet, P.M.; Reedijk, J.; Brabec, V.; Correlation between cytotoxicity and DNA binding of polypyridyl ruthenium complexes; Biochemistry, 1995, 34, 12369–12378. 29. Karidi, K.; Garoufis, A.; Hadjiliadis, N.; Reedijk, J.; Solid-phase synthesis, characterization and DNA binding properties of the first chloro(polypyridyl)ruthenium conjugated peptide complex; J. Chem. Soc., Dalton Trans., 2005, 728–734. 30. Lopes, L.G.F.; Wieraszko, A.; El-Sherif, Y.; Clarke, M.J.; The trans-labilization of nitric oxide in RuII complexes by C-bound imidazoles; Inorg. Chim. Acta, 2001, 312, 15–22. 31. Lang, D.R.; Davis, J.A.; Lopes, L.G.F.; Ferro, A.A.; Vasconcellos, L.C.G.; Franco, D.W.; Tfouni, E.; Wieraszko, A.; Clarke, M.J.; A controlled NO-releasing compound: synthesis, molecular structure, spectroscopy, electrochemistry, and chemical reactivity of R,R,S,Strans-[RuCl(NO)(cyclam)]2+(1,4,8,11-tetraazacyclotetradecane); Inorg. Chem., 2000, 39, 2294–2300. 32. Karidi, K.; Garoufis, A.; Tsipis, A.; Hadjiliadis, N.; den Dulk, H.; Reedijk, J.; Synthesis, characterization, in vitro antitumor activity, DNA-binding properties and electronic structure (DFT) of the new complex cis-(Cl,Cl)[RuIICl2(NO+)(terpy)]Cl; J. Chem. Soc., Dalton Trans., 2005, 1176–1187. 33. Slocik, J.M.; Shepherd, R.E.; Coordination of Ru(NO)Cl3 to the tripeptides gly–gly–gly and gly–gly–his: N-terminal amine–amide and C-terminal imidazole–amide functionalities in bidentate chelation; Inorg. Chim. Acta, 2000, 311, 80–94. 34. Patra, A.K.; Mascharak, P.K.; A ruthenium nitrosyl that rapidly delivers NO to proteins in aqueous solution upon short exposure to UV light; Inorg. Chem., 2003, 42, 7363– 7365. 35. Bordini, J.; Hughes, D.L.; Neto, J.D.D.; Da Cunha, C.J.; Nitric oxide photorelease from ruthenium salen complexes in aqueous and organic solutions; Inorg. Chem., 2002, 41, 5410–5416. 36. Karidi, K.; Garoufis, A.; Hadjiliadis, N.; Lutz, M.; Spek, A.L.; Reedijk, J.; Synthesis, characterization, and DNA-binding studies of nitro(oligopyridine)ruthenium(II) complexes; Inorg. Chem., 2006, 45, 10282–10292. 37. Cardo, L.; Hannon, M.J.; Design and DNA-binding of metallo-supramolecular cylinders conjugated to peptides; Inorg. Chim. Acta, 2008, doi; 10.1016/j.ica.2008.02.050. 38. (a) Hannon, M.J.; Moreno, V.; Prieto, M.J.; Molderheim, E.; Sletten, E.; Meistermann, I.; Isaac, C.J.; Sanders, K.J.; Rodger, A.; Intramolecular DNA coiling mediated by a metallo supramolecular cylinder; Angew. Chem., Intl. Ed., 2001, 40, 879–884. (b) Uerpmann, C.; Malina, J.; Pascu, M.; Clarkson, G.J.; Moreno, V.; Rodger, A.; Grandas, A.; Hannon, M. J.; Design and DNA binding of an extended triple-stranded metallo-supramolecular cylinder; Chem., Eur. J., 2005, 11, 1750–1756. (c) Meistermann, I.; Moreno, V.; Prieto, M.J.; Moldrheim, E.; Sletten, E.; Khalid, S.; Rodger, P.M.; Peberdy, J.C.; Isaac, C.J.; Rodger, A.,

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DNA Recognition and Binding by Peptide–Metal Complex Conjugates Hannon, M.J.; Intramolecular DNA coiling mediated by metallo-supramolecular cylinders: differential binding of P and M helical enantiomers; Proc. Natl. Acad. Sci. USA., 2002, 99, 5069–5074. (a) Cerasino, L.; Hannon, M.J.; Sletten, E.; DNA three-way junction with a dinuclear iron(II) supramolecular helicate at the center: A NMR structural Study; Inorg. Chem., 2007, 46, 6245–6251. (b) Malina, J.; Hannon, M.J.; Brabec, V.; Recognition of DNA threeway junctions by metallosupramolecular cylinders: gel electrophoresis studies; Chem. Eur. J., 2007, 13, 3871–3877. (c) Childs, L.J.; Malina, J.; Rolfsnes, B.E.; Pascu, M.; Prieto, M.J.; Broome, M.J.; Rodger, P.M.; Sletten, E.; Moreno, V.; Rodger, A.; Hannon, M.J.; A DNA-binding copper(I) metallo-supramolecular cylinder that can act as an artificial nuclease; Chem. Eur. J., 2006, 12, 4919–4927. Brunner, J.; Barton, J.K.; Targeting DNA mismatches with rhodium intercalators functionalized with a cell-penetrating peptide; Biochemistry, 2006, 45, 12295–12302. Wender, P.A.; Mitchell, D.J.; Pattabiraman, K.; Pelkey, E.T.; Steinman, L.; Rothbard, J.B.; The design, synthesis, and evaluation of molecules that enable or enhance cellular uptake: Peptoid molecular transporters; Proc. Natl. Acad. Sci. USA, 2000, 97, 13003–13008. Jackson, B.A.; Alekseyev, V.Y.; Barton, J.K.; A versatile mismatch recognition agent: specific cleavage of a plasmid DNA at a single base mispair; Biochemistry, 1999, 38, 4655–4662. Copeland, K.D.; Lueras, A.M.K.; Stemp, E.D.A.; Barton, J.K. DNA cross-linking with metallointercalator-peptide conjugates; Biochemistry, 2002, 41, 12785–12797. Lasey, R.C.; Banerji, S.S.; Ogawa, M.Y.; Synthesis and characterization of a sequencespecific DNA-binding protein that contains ruthenium polypyridyl centers; Inorg. Chim. Acta, 2000, 300–302, 822–828. Hodges, R.S.; De novo design of alpha-helical proteins: basic research to medical applications; Biochem. Cell Biol., 1996, 74, 133–154. Sardesai, N.Y.; Zimmermann, K.; Barton, J.K.; DNA recognition by peptide complexes of rhodium(III): Example of a glutamate switch; J. Am. Chem. Soc., 1994, 116, 7502–7508. Hastings, C.A.; Barton, J.K.; Perturbing the DNA sequence selectivity of metallointercalator-peptide conjugates by single amino acid modification; Biochemistry, 1999, 38, 10042–10051. Sardesai, N.Y.; Barton, J.K.; DNA recognition by metal-peptide complexes containing the recognition helix of the phage 434 repressor; J. Biol. Inorg. Chem., 1997, 2, 762– 771. Myari, A.; Hadjiliadis, N.; Garoufis, A.; Synthesis and characterization of the diastereomers L- and D-[Ru(bpy)2(m-bpy-L-Arg-Gly-L-Asn-L-Ala-L-His-L-Glu-LArg)]Cl2.1H NMR studies on their interactions with the deoxynucleotide duplex d[(5′GCGCTTAAGCGC-3′)2] and d[(5′-CGCGATCGCG-3′)2]; J. Inorg. Biochem., 2005, 99, 616–626. Deibert, M.; Grazulis, S.; Janulaitis, A.; Siksnys, V.; Huber, R.; Crystal structure of MunI restriction endonuclease in complex with cognate DNA at 1.7 Å resolution; EMBO J., 1999, 18, 5805–5816. Barton, J.K.; Metals and DNA: Molecular left-handed complements; Science, 1986, 233, 727–734. Fitzsimons, M.P.; Barton, J.K.; Design of a synthetic nuclease: DNA hydrolysis by a zinc-binding peptide tethered to a rhodium intercalator; J. Am. Chem. Soc., 1997, 119, 3379–3380. Houser, R.P.; Fitzsimons, M.P.; Barton, J.K.; Metal-dependent intramolecular chiral induction: The Zn2+ complex of an ethidium-peptide conjugate; Inorg. Chem., 1999, 38, 1368–1370. Copeland, K.D.; Fitzsimons, M.P.; Houser, R.P.; Barton, J.K.; DNA hydrolysis and Oxidative Cleavage by Metal-Binding Peptides Tethered to Rhodium intercalators; Biochemistry, 2002, 41, 343–356.

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55. Pyle, A.M.; Rehmann, J.P.; Meshoyrer, R.; Kumar, C.V.; Turro, N.J.; Barton, J.K.; Mixedligand complexes of ruthenium(II): Factors governing binding to DNA; J. Am. Chem. Soc., 1989, 111, 3051–3058. 56. (a) Liang, Q.; Eason, P.D.; Long, E.C.; Metallopeptide-DNA interactions: Site-selectivity based on amino acid composition and chirality; J. Am. Chem. Soc., 1995, 117, 9625–9631. (b) Huang, X.; Pieczko, M.E.; Long, E.C.; Combinatorial optimization of the DNA cleaving Ni(II)·Xaa-Xaa-His metallotripeptide domain; Biochemistry, 1999, 38, 2160–2166. (c) Fang, Y.-Y.; Ray, B.D.; Claussen, C.A.; Lipkowitz, K.B.; Long, E.C.; Ni(II)·Arg-Gly-HisDNA interactions: Investigation into the basis for minor-groove binding and recognition; J. Am. Chem. Soc., 2004, 126, 5403–5412. (d) Fang, Y.-Y.; Claussen, C.A.; Lipkowitz, K.B.; Long, E.C.; Diastereoselective DNA cleavage recognition by Ni(II)·Gly-Gly-Hisderived metallopeptides; J. Am. Chem. Soc., 2006, 128, 3198–3207. (e) Jin, Y.; Lewis, M.A.; Gokhale, N.H.; Long, E.C.; Cowan, J.A.; Influence of stereochemistry and redox potentials on the single- and double-strand DNA cleavage efficiency of Cu(II) and Ni(II)·LysGly-His-derived ATCUN metallopeptides; J. Am. Chem. Soc., 2007, 129, 8353–8361. 57. (a) Ananias, D.C.; Long E.C.; DNA strand scission by dioxygen + light-activated cobalt metallopeptides; Inorg. Chem., 1997, 36, 2469–2471. (b) Ananias, D.C.; Long E.C.; Highly selective DNA modification by ambient O2-activated Co(II)·Lys-Gly-His metallopeptides; J. Am. Chem. Soc., 2000, 122, 10460–10461. 58. Sissi, C.; Rossi, P.; Felluga, F.; Formaggio, F.; Palumbo, M.; Tecilla, P.; Toniolo, C.; Scrimin, P.; Dinuclear Zn2+ complexes of synthetic heptapeptides as artificial nucleases; J. Am. Chem. Soc., 2001, 123, 3169–3170. 59. Shullenberger, D.F.; Eason, P.D.; Long, E.C.; Design and synthesis of a versatile DNAcleaving metallopeptide structural domain; J. Am. Chem. Soc., 1993, 115, 11038–11039. 60. (a) Welch, J.T.; Sirish, M.; Lindstrom, K.M.; Franklin, S.J.; De novo nucleases based on HTH and EF-hand chimeras; Inorg. Chem., 2001, 40, 1982–1984. (b) Sirish, M.; Franklin, S.J.; Hydrolytically active Eu(III) and Ce(IV) EF-hand peptides; J. Inorg. Biochem., 2002, 91, 253–258.

13 Artificial Restriction Agents: Hydrolytic Agents for DNA Cleavage Fabrizio Mancin and Paolo Tecilla

13.1 Introduction The inertness of DNA toward hydrolytic cleavage is tremendous. Indeed, the estimated half-life times for a single P–O bond cleavage at 25 °C and pH 7 range from hundreds of thousands to hundreds of millions of years.1 Such hydrolytic resistance contributes to the preservation of the genetic information, but also represents a serious obstacle, since several biological processes involving DNA manipulation, e.g. expression and duplication, repair of damages and elimination of foreign DNA, require hydrolytic reactions. The solution of this problem is provided by a wealth of hydrolytic enzymes, nucleases and topoisomerases, which efficiently catalyse DNA scission. Many such enzymes contain metal ions in their active sites, mainly Ca(II), Mg(II) and Zn(II), that play a fundamental role in their catalytic action.1b,2 As a matter of fact, the ability of metal ions to promote the hydrolytic cleavage of esters has been well known by chemists for decades. However, the development of synthetic agents capable to accelerate the hydrolysis of DNA has attracted substantial research efforts only in the last 20 years.2a,3 There are several reasons that justify the interest in such systems. First, the mechanistic information obtained in the study of artificial agents could lead to a better understanding of the chemistry of the corresponding hydrolytic enzymes. Second, they could be employed for detoxification of pesticides and chemical weapons, which often have phosphate Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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ester-like structures. Third, efficient hydrolytic agents for phosphate diesters could be employed as artificial restriction enzymes for molecular biology. Such systems would be highly valuable because they could, in principle, be designed to cleave DNA with sequence selectivity different from that of the natural enzymes. The latter, in fact, recognize DNA sequences of only 4, 6 or 8 bases, and such selectivity may be inadequate for the precise control of the manipulation of large DNA of higher species. Finally, the realization of antiDNA drugs can be envisaged in a more distant future. Since the first example of an artificial hydrolitic agent for DNA cleavage, reported by J. K. Barton in 1987,4 several promising systems have been proposed up to the description of the first studies on DNA manipulation with artificial agents, reported in very recent years by Komiyama and coworkers.5 This chapter highlights the progress reported during these two decades toward obtaining synthetic nucleases. Particular attention will be devoted to the strategies that can be pursued to increase the efficiency and the sequence selectivity of the systems.

13.2 DNA Hydrolysis Information available on the mechanism of spontaneous hydrolysis of DNA is obviously scarce due to its virtually absent reactivity. The commonly accepted pathway for the hydrolysis of deoxynucleotide phosphates involves the nucleophilic attack of a water (or hydroxide) oxygen at the phosphorus to give a five-coordinate phosphate intermediate or transition state (Figure 13.1).3o,p,6 Subsequent cleavage of either the P-O3′ or P-O5′ (usually P-O3′ cleavage in the enzymatic process) causes a strand scission, yielding the R–OH and R–O–PO3(H2) termini. As mentioned before, such a reaction is exceedingly slow in physiological conditions (i.e, aqueous solution at 25 °C and pH 7) and reaction rate constants of about 1 × 10−15 s−1 have

Figure 13.1 Reaction pathway for DNA hydrolysis. The enzymatically promoted P-03′ scission is shown

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been estimated.1a As a consequence, spontaneous DNA degradation occurs mainly through different pathways, involving either C–O cleavage, nucleobase ring opening or radical attack (Figure 13.2).1a Metal ions can assist the P–O hydrolytic cleavage of phosphodiesters in different ways (Figure 13.3). Acting as Lewis acids, they can activate the phosphate group toward the attack of the nucleophile, increase the leaving group ability of the departing alcohol and activate a metal-coordinated water molecule as a nucleophile.1b Moreover, indirect activation modes are possible, with metal bound hydroxides or water molecules performing general base or acid catalysis. As observed in many enzymatic processes, several metal centers may act cooperatively to accelerate these mechanistic steps. Accordingly, multinuclear complexes are often more active than their mononuclear counterparts.1b The exceptional high resistance of DNA toward hydrolysis makes even a simple kinetic investigation of the cleavage process quite challenging. However, supercoiled plasmid DNA is a more accessible substrate and, because of this, very popular

Figure 13.2 Alternative DNA degradation pathways

Figure 13.3 Possible activation modes provided by metal ions for acceleration of the hydrolysis of phosphate diesters

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Figure 13.4 The three forms of plasmid DNA

for such studies.3r This particular form of DNA, which is commonly found in bacteria, is a cyclic supercoiled double strand made by several thousands of base pairs (Figure 13.4). One single-strand scission unravels the supercoiled DNA (form I) to a relaxed circular one (nicked, form II), while a second scission on the complementary strand, within about twelve base pairs from the first one, generates a linear DNA form (form III). These three DNA forms can be easily separated and quantified by gel electrophoresis, thus allowing a simple and rapid kinetic analysis. Moreover, supercoiled DNA is somehow more reactive than a short linear DNA fragment and this makes the study of the reaction easier, even in the presence of relatively poor catalysts. Such an increased reactivity is due: (a) to the obvious reason that the cleavage of a single phosphodiester bond in a large DNA molecule is statistically more likely than in a smaller DNA fragment and (b) to the internal strain resulting in ground-state destabilization.3r Finally, two other peculiar and relevant features of plasmid DNA are: (i) the possibility to discriminate between a single-strand or a double-strand cleavage simply by a statistical analysis of the relative abundance of forms II and III produced in the reaction; (ii) the possibility to enzymatically religate form III or form II giving both a closed circular form and concatemer DNA which can be easily detected by gel electrophoresis.3r Religation experiments are important in the mechanistic investigation of DNA cleavage because only fragments that derive from a hydrolytic process can be religated by the enzyme. Therefore, the success or the failure of this experiment may give important support to or seriously question the occurrence of a clean hydrolytic process. A careful mechanistic investigation becomes extremely important when metal ions with known redox chemistry are used for the development of artificial nucleases. For example, it is well known that Cu(II) and Fe(III) complexes, in the proper conditions, cleave DNA by an oxidative pathway.7 This process is so efficient that it always competes with the hydrolytic cleavage. Because the reactivity of oxidative agents depends usually on cofactors such as H2O2, molecular oxygen and reducing agents, and may produce diffusible radicals, the absence of externally added cofactors, the insensitivity to radical scavengers and the preservation of reactivity under anaerobic reactivity are generally considered sufficient evidence of a hydrolytic

Free Ions and Mononuclear Complexes 373

Figure 13.5 The [Zn2(pyrimol)2]2+ complex: the first Zn-based artificial oxidative nuclease

reaction. However, oxidative agents that cleave DNA through nondiffusible radicals have been known for a long time and, more recently, several complexes able to perform oxidative cleavage in the absence of coreactants and in anaerobic conditions have been reported.8 Remarkably, in a very recent example by Reedijk and coworkers,9 such behaviour has been described for the first time, even in the case of Zn(II) complexes, a metal ion so far considered only capable of promoting hydrolytic chemistry. This peculiar reactivity is probably related to the pirymol ligand employed (Figure 13.5), but highlights the importance of a carefully performed mechanistic investigation and indicates that any claim of hydrolytic reactivity, particularly in the case of Cu(II) or iron complexes, not supported by clear-cut evidence, such as enzymatic religation or fragments identification, should be considered with caution.

13.3 Free Ions and Mononuclear Complexes At the beginning of the last decade it was demonstrated that aqueous trivalent lanthanide ions accelerate the hydrolysis of DNA.10 Subsequent papers by Schneider and coworkers reported a systematic investigation of such ions aimed at determining the relevant kinetic parameters for the nicking of plasmid DNA.11 The different ions show similar maximum reactivities under saturation conditions (kmax 0.7–1.0 × 10−4 s−1, 37 °C, pH 7.0), but different affinities for DNA. The latter increase along the series from La(III) (Ka = 5.0 × 102 M−1) to Er(III) (Ka = 1.7 × 104 M−1). The combination of moderate reactivity with low affinity for the substrate is such that substantial degradation of DNA can be obtained only at high metal ion concentrations. In contrast, late lanthanides, such as Tm(III), Yb(III) and Lu(III), display a remarkable reactivity, even at low concentrations, due to their high affinities for DNA. However, a decrease of the cleavage rate is observed at high metal ion concentrations, probably as a consequence of their aggregation, with formation of less active species. Among lanthanides, cerium is peculiar in its ability to reach a stable tetravalent oxidation state. Ce(IV) is much more efficient in hydrolysing DNA than the

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trivalent lanthanide ions. At pH 7.0 and 50 °C, the half-life of the dinucleotide TpT is reduced to 3.6 hours, which amounts to more than 11 orders of magnitude rate acceleration over the spontaneous hydrolysis reaction.10a,12 As a result, Ce(IV) is 20 to 1000 times more efficient than any other trivalent lanthanide, with comparable high efficacy toward single- and double-stranded linear DNA.13 The source of this remarkable reactivity has been attributed by Komiyama and coworkers to the great electron-withdrawing ability of this tetravalent ion.3m Surprisingly, free Ce(IV) ions are inactive in promoting plasmid DNA hydrolysis, probably due to steric hindrance between the Ce(IV) hydroxide gels, which form at neutral pH, and the substrate.14 In fact, when converted into homogeneous solutions by the addition of solubilizing agents (dextran, polyvinylpirrolidone), Ce(IV) recovers activity toward plasmid DNA and is about ten times more reactive than trivalent lanthanide ions.15 Aqueous lanthanide ions are toxic for biological systems and, more generally, free metal ions are not suitable candidates for preparing appealing catalysts. The formation of stable complexes with proper ligands is therefore mandatory. However, this task turns out to be quite difficult in the case of lanthanide ions for two reasons: (i) they undergo very facile ligand exchange; (ii) the catalytic activity requires an unsaturated coordination sphere to allow interaction with water (nucleophile) and the substrate. Stable complexes can be obtained by using polyaminocarboxylate derivatives, crown ethers or azacrown ligands.3i In many cases, however, the formation of the complex results in a lower reactivity than that observed with the free metal ions, particularly in the case of polyaminocarboxylate complexes, due to the decrease in the overall charge of the complex and to the saturation of the metal ion coordination sphere. Neutral ligands, such as azacrowns or crown ethers, are thus more suitable, in order to retain the reactivity of the free metal ion, although their coordination strength is not so high and the complexing agent must be used in large excess to ensure complete formation of soluble complexes.16 In the case of Ce(IV), the reactivity decrease brought about by the use of anionic ligands leads to an interesting substrate selectivity. In fact, the Ce(IV)EDTA complex is able to cleave single-stranded DNA (ssDNA), but not doublestranded DNA (dsDNA).17 The reaction is slow, but highly selective and, as a consequence, gap sites or bulges present in a dsDNA that expose short sequences of ssDNA are cleaved without touching the rest of the macromolecule.18 Such behaviour is the basis of the ARCUT system proposed by Komiyama and coworkers,5 which will be described in detail later. Metal ions other than lanthanides are scarcely active in promoting DNA hydrolysis. Nevertheless, a few mononuclear metal complexes have been reported to act as artificial nucleases. The Co(III)-TAMEN complex (Figure 13.6) cleaves plasmid DNA, at 37 °C and pH 7.6, with a first-order rate constant of 5 × 10−5 s−1.19 Although the concentration of complex required, 1mM, is relatively high, suggesting a poor interaction with the substrate, the mechanism is fully hydrolytic, as the nicked DNA can be religated. This complex is also active toward single-stranded DNA. Zn(II) is one of the metal ions most frequently found in hydrolytic metalloenzymes, but the activities reported for simple mononuclear complexes are usually low. One remarkable exception is the Zn(II)-BHTDE complex (BHDTE = N,N′-bis(benzylhistidyl)diethylenetriamine, Figure 13.7) reported by Ichikawa and

Free Ions and Mononuclear Complexes 375

Figure 13.6 The TAMEN ligand

Figure 13.7 [Zn(H2BHTDE)OH]3+ and [Zn(quercetin)2] complexes

coworkers,20 which is quite reactive toward plasmid DNA: at pH 7, 50 °C and 3 mM complex concentration, the supercoiled form of plasmid DNA is completely cleaved after 24 h, leading to both nicked and linear products. From this data a rate constant of 4 × 10−5 s−1 can be roughly estimated. The cleavage efficiency shows a maximum at pH 6.5–7.0, suggesting the possibility that the protonated primary amino groups play a role in increasing DNA affinity. Another interesting example is the [Zn(quercetin)2] complex,21a (Figure 13.7), which at pH 7.2, 37 °C and 100 mM complex concentration cleaves supercoiled plasmid DNA with a rate constant of 1.7 × 10−4 s−1, which corresponds to a half-life of about 1 hour. Linearized DNA could be religated with the enzyme T4 ligase, ruling out the possibility of oxidative mechanisms involving phenoxyl radicals and confirming a pure hydrolytic cleavage. Remarkably, the similar Mn(II)-quercetin complex also shows hydrolytic activity very close to that of the Zn(II) complex (k = 1.3 × 10−4 s−1, at pH 7.2, 37 °C and 100 mM complex concentration).21b Because of their redox properties, Cu(II) complexes have been frequently used in the development of agents for the oxidative cleavage of DNA,7 but there are also several examples of Cu(II) complexes reported to cleave DNA with a hydrolytic mechanism. Burstyn and coworkers published the first example of a Cu(II)-based hydrolytic agent in 1996.22a It was shown that at a 25 mM concentration the Cu(II)-TACN complex (Figure 13.8) cleaves supercoiled DNA at 50 °C and pH 7.8 with an

376

Artificial Restriction Agents

Figure 13.8 Structures of some polyamine ligands used for the formation of mononuclear Cu(II) complexes active in DNA cleavage

estimated rate constant of ca. 1.5 × 10−5 s−1. A decrease of reactivity was observed at higher complex concentrations probably due to the formation of unreactive mhydroxo dimers.22b The complex is also active in the degradation of single-stranded DNA. However, anaerobic experiments showed that the activity in the absence of oxygen is reduced by 30%, indicating at least the simultaneous occurrence of both hydrolytic cleavage and oxidative cleavage. Later on, Fujii and coworkers reported that the Cu(II)-TACH complex (Figure 13.8) promotes the oxygen-independent cleavage of plasmid DNA.23a Saturation kinetics were observed with a kmax value of 1.2 × 10−3 s−1 and a Ka value of 1.3 × 104 M−1 at 35 °C and pH 8.1. Such reactivity is among the highest reported so far and corresponds to a half-life time for the supercoiled form of 20 minutes in the presence of 75 mM Cu(II) complex. Several other complexes of macrocyclic and linear triamine ligands were tested and found to be scarcely reactive. On the basis of these results and of the relatively good affinity of the Cu(II)-TACH complex for DNA, the authors suggest the possibility of specific binding of the catalyst to the substrate. A detailed analysis of complex speciation and pH reactivity profiles indicate that the monohydroxo complex is the active species at pH values above neutrality, as observed in the hydrolytic cleavage of model phosphate diesters.23b However, linearized plasmid DNA could not be enzymatically religated.23b Similar results have also been obtained for the related Cu(II)-TACI complex (Figure 13.8), which again shows an oxygen-independent activity (k = 2.3 × 10−3 s−1 at 48 mM complex concentration), but fails the enzymatic religation test.24 A pure hydrolytic mechanism, demonstrated both by oxygen-independent activity and enzymatic religation, characterizes the activity of the Cu(II) complexes of natural aminoglycosides, such as neamine (Figure 13.8) studied by Cowan and coworkers.25 With the Cu(II)-neamine complex, saturation kinetics were observed with a kmax value of 5.2 × 10−4 s−1 and a Ka value of 2.4 × 105 M−1 at 37 °C and pH 7.3. The very high affinity of the complex for DNA is attributed to the tight binding of the positively charged aminoglycoside ligand to DNA; this apparently makes the system very effective even at low concentrations, reducing the half-life time for the

Bimetallic Complexes

377

supercoiled form, in the presence of 25 mM complex concentration, to only 26 minutes. In the last few years several other reports on copper complexes able to cleave DNA have appeared in the literature.26 In most cases, the authors suggest a hydrolytic mechanism, basing their proposal only on the observation of reactivity in anaerobic conditions and on insensitivity to radical scavengers. However, although these behaviours are compatible with a hydrolytic mechanism, they have also been reported in the case of oxidative systems (see Section 13.2) and, therefore, they cannot be taken as clear-cut mechanistic evidence. Particularly when a metal ion such as Cu(II) is involved, the elucidation of the mechanism requires more stringent evidence, such as, for example, enzymatic religation or chemical identification of the cleaved DNA fragments. In the absence of such experiments the final decision on the true nature of the DNA cleavage remains uncertain.

13.4 Bimetallic Complexes The extraordinary catalytic efficiency of natural metallonucleases most often relies on the cooperative action of two or more metal ions. Available X-ray structures of these proteins indicate that most of these metal ions are placed within a narrow range of distances from each other.2 Accordingly, in a synthetic catalyst a precise spatial localization of the ions appears to be mandatory to ensure multiple interactions with the substrate and, consequently, to take advantage simultaneously of all the different activation modes that metal ions can provide for the hydrolysis of phosphate esters (substrate, leaving group and nucleophile activation).1b On the basis of these guidelines, a series of bimetallic complexes have been synthesized and investigated as catalysts for the hydrolytic cleavage of DNA. In 1996, Schneider and coworkers studied a 30-membered azacrown macrocycle (Figure 13.9a) that can bind two Eu(III) or Pr(III) ions.27 The Pr(III) complex was able to promote plasmid DNA nicking with a kmax = 2.8 × 10−4 s−1 and Ka = 3.0 × 103 M−1 at 37 °C and pH 7.0. However, the activity of the binuclear complex was only twofold that of the free metal ion. More effective are the Er(III)2 complexes of a Schiff-base-containing macrocycle (Figure 13.9b), proposed by Zhu and coworkers, which degrades supercoiled DNA at 37 °C, pH 7.0 with a maximum rate constant of 1.0 × 10−3 s−1.28 Unfortunately

Figure 13.9 Macrocyclic ligands capable of binding two lanthanide ions reported by Schneider (a) and Zhu (b)

378

Artificial Restriction Agents

Figure 13.10 The HXTA ligand

such high reactivity is counterbalanced by a low affinity for the substrate (Ka = 1.0 × 103 M−1) so that high concentrations of complex are required to obtain high reaction rates. A very efficient dicerium complex of the polyaminocarboxylate ligand HXTA (Figure 13.10) has been reported by Que and coworkers.29 At a concentration of 10 mM, pH 8 and 37 °C, the Ce2-HXTA complex cleaves plasmid DNA with a rate constant of 1.4 × 10−4 s−1, which corresponds to a half-life of 1.4 hours for the nicking process. Interestingly, double-strand scission is preferred to single-strand scission and the complex also cleaves linear double-stranded DNA with high regioselectivity (more than 90%) favouring the scission of the P-O3′ bond. Fe(III) is present in the active site of some phosphatases, but it has been rarely employed in artificial nucleases. Interestingly, the few Fe(III) complexes reported in the literature with hydrolytic activity in DNA cleavage are all binuclear. The first example, described by Schnaith, Que and coworkers in 1994,30 is based on a tetrabenzimidazolylmethyl derivative of 1,3-diamino-2-hydroxypropane (Figure 13.11, HPTB) and shows a surprising reactivity: the Fe(III)2-HPTB complex requires ‘oxidative’ conditions, i.e. the presence of H2O2 or O2 and a reductant (dithiothreitol or ascorbate), but yields DNA fragments that are consistent with hydrolytic cleavage. In fact, the linearized plasmid DNA can be enzymatically religated with a quantitative conversion, indicating that only the products of P-O3′ bond scissions are formed. This high regioselectivity is also supported by 3′- and 5′-end labelling studies. The cleavage of plasmid DNA is so efficient that, in the presence of 10 mM complex, at 25 °C and pH 8.0, the supercoiled form is completely degraded immediately after mixing. The extent of cleavage depends on the concentration of the complex, but, surprisingly, it does not increase with extended reaction times. To justify this unusual reactivity, the authors proposed a mechanism involving nucleophilic attack of a metal-ion-bound peroxide to the phosphodiester bond.31 Similar behaviour has been recently reported for the dinuclear complex [CpFe(CO)2]2 which hydrolytically cleaves DNA at very low concentrations in the presence of H2O2 or other peroxides.32 Another, again very efficient, diiron(III) complex, based on a benzimidazolylmethyl derivative of 1,4,7-triazaheptane (Figure 13.11, DTPB), has been investigated by Liu and coworkers.33 In the presence of 100 mM of Fe(III)2-DTPB complex, at 37 °C and pH 7.0, the supercoiled DNA is degraded with a rate constant of 2.1 × 10−3 s−1, which corresponds to a half-life of 5 minutes. This is the current ‘world record’ for the cleavage of plasmid DNA under ‘hydrolytic conditions’. The extent of cleavage does not depend on the presence of O2 and the linearized DNA can be

Bimetallic Complexes

Figure 13.11 (DTPB)

379

Bimetallic ligand for Fe(III) reported by Schnaith and Que (HPTB), and Liu

quantitatively religated, thus confirming the occurrence of a hydrolytic mechanism with the sole cleavage of the P-O3′ bond. The complex shows a remarkable affinity for plasmid DNA (Ka = 1 × 105 M−1), probably related to the high positive charge of its bimetallic structure. The comparison with other bimetallic complexes indicates that a m-oxo bridge between the two metal ions and the presence of a labile BF4− ligand play a key role in determining the reactivity: the first by ensuring the proper intermetallic distance and the second by allowing a facile substitution to yield free coordination sites for interaction with the substrate. Surprisingly, examples of bimetallic systems based on Cu(II) or Zn(II) are scarce and show a disappointingly low reactivity.34 The only important exception is a binuclear Zn(II)-binding heptapeptide (Figure 13.12), described by Scrimin and coworkers,35 which cleaves plasmid DNA with a first-order rate constant of 1.0 × 10−5 s−1 at pH 7.0, 37 °C and 3.6 mM complex concentration. Due to the presence of several a-disubstituted amino acids, the peptide is folded to a relevant extent in a 310-helical conformation, and the two metal chelating TACN moieties face each other at a distance of about 6 Å (the pitch of the 310-helix). Even if such distance is larger than that found in bimetallic hydrolytic enzymes, the system is about 20 times more reactive than its monometallic counterpart and the cooperativity between the two metal centres has been clearly demonstrated with Zn2+-concentration-dependent experiments. The high cooperativity observed has been justified by assuming the formation of a sandwich-like complex in which the two Zn(II)-TACN complexes interact with a single DNA phosphate group, taking full advantage of their complementary roles for the hydrolytic cleavage.

380

Artificial Restriction Agents

Figure 13.12

Chemical structure of the Scrimin’s ditopic Zn(II)-binding heptapeptide

An impressive high reactivity was recently reported by Yu and coworkers,36a who studied dimetallic Zn(II) complexes made by linking two cyclen ligands with different spacers bearing alcoholic, phenolic or pyridinic moieties. The authors claim complete nicking of plasmid DNA in only five minutes at pH 7.1, 37 °C and 25 mM complex concentration. Longer times of incubation result in further fragmentation and DNA smearing on the electrophoresis gel. However, the complexes are active only in the presence of vitamin C as reducing agent. Therefore, the occurrence of some concurrent oxidative degradation pathway, involving the ligand backbone or adventitious redox active metal ions, is possible. The same type of reactivity was also observed in the case of the Zn(II) complexes of uracil-cyclen conjugate ligands.36b These results, although impressive if the mechanism proves to be really hydrolytic, stress the importance of using caution when studying the mechanism of DNA cleavage, even when Zn(II) is employed as metal ion. Indeed this natural substrate is really stable toward hydrolysis, but is also very delicate from other points of view and competitive cleavage pathways are always possible.

13.5 Conjugation with DNA-affine Subunits Simple considerations, supported by several examples reported in the case of oxidative DNA cleaving agents,37 suggest that the reactivity of hydrolytic systems should be strongly enhanced when their structures comprise groups with high DNA affinity. Furthermore, such groups could also provide sequence specificity to the cleaving agents and thus open the way toward the obtainment of real artificial restriction enzymes. The obvious choice among the different families of compounds able to increase DNA affinity is the utilization of intercalators. Quite surprisingly, examples of metal complexes appended to intercalating groups as hydrolytic agents are rare, and the effects of such elements on the reactivity of the systems are not always straightforward. For example, in the case of the Scrimin’s peptide discussed above,35 the introduction of an acridine moiety at the N-terminus of the heptapeptide resulted in a slightly higher activity at low catalyst concentration compensated by a decrease at higher concentrations.

Conjugation with DNA-affine Subunits 381

Figure 13.13 Artificial nucleases based on ruthenium and rhodium intercalators reported by Burton and coworkers

In 1987, Barton and coworkers published the first example of an artificial system with DNA hydrolytic cleavage activity.4 It was based on a ruthenium intercalator with two arms having the role of metal-binding moieties (Figure 13.13a). The Zn(II) and the Cd(II) binuclear complexes of this ligand cleaved plasmid DNA with high efficiency. At 37 °C and pH 8.5, 40% of the supercoiled form was degraded in the presence of a 7 mM concentration of the complex after 5 hours of incubation (a first-order rate constant of 3 × 10−5 s−1 can be roughly estimated). Religation experiments showed that the hydrolysis occured randomly at the P-O3′ and P-O5′ bonds. However, the affinity of the ligand for the metal ions seemed to be quite low and a large excess of metal ion was required. Unfortunately, no evidence concerning the role played by the intercalator was reported. Later on, Barton and coworkers studied the reactivity of a mononuclear Zn(II)binding peptide, tethered to a rhodium complex as major groove intercalator (Figure 13.13b).38 The system promotes plasmid DNA cleavage with a rate constant of 2.5 × 10−5 s−1 at pH 6, 37 °C and 5 mM complex concentration, and shows similar activity toward linear double-stranded DNA. Analysis of the fragments produced showed that the cleavage occurs only at the P-O3′ bonds with a modest sequence selectivity for 5′-Pu-Py-Pu-Py-3′sites (with cleavage at Py), presumably the result of the Rh complex binding selectivity. The reactivity decreases as the pH approaches neutrality. The peptide is scarcely structured and this is probably the source of the weak binding of the Zn(II) ion. Although the presence of the rhodium complex intercalator is of paramount importance for the activity of the complex, its substitution with a different rhodium-based intercalator, which preferentially binds at mismatches, or with the ethidium intercalator led to unreactive systems.38b Schneider and coworkers appended two naphthalene groups to an azacrown ligand via C6 alkyl spacers (Figure 13.14a) and used it as a cofactor for Eu(III)promoted DNA cleavage.39 The results are puzzling because conjugation with the intercalating units led to an increase in the intrinsic reactivity with regard to the

382

Artificial Restriction Agents

Figure 13.14 Intercalator-metal binding unit conjugates reported by Schneider (a), Nakamura (b) and Tonellato (c)

free metal ion, but to a decrease in the DNA affinity. Finally, an earlier report by Nakamura and Hashimoto,40 which investigated the reactivity of a hydroxamic acid linked to a phenanthridine intercalator (Figure 13.14b) in the presence of different lanthanide ions, points attention to the length of the spacer that tethers the intercalating unit to the catalytic group as a key element for cleavage activity. This result is confirmed by the reactivity of a series of Zn(II)-cis-cis-triaminocyclohexane complex–anthraquinone intercalator conjugates, linked by alkyl spacers of different length, reported by Tonellato and coworkers (Figure 13.14c).41 At a concentration of 5 mM, the complex of the derivative with a C8 alkyl spacer cleaves supercoiled DNA with a rate of 4.6 × 10−6 s−1 at pH 7 and 37 °C. Saturation kinetics have been observed with a binding constant (Ka) of about 1 × 104 M−1, in agreement with the reported DNA affinity of anthraquinone. Thus, the conjugation of the metal complex with the intercalating group led to a 15-fold increase in the cleavage efficiency when compared with the Zn-triaminocyclohexane complex lacking the anthraquinone moiety. Comparison of the reactivity of the different complexes showed a remarkable increase in DNA cleaving efficiency due to the increase in the spacer length. In the case of the shortest spacer (C4) no cleavage was observed, indicating that the advantages derived by the increased DNA affinity may be cancelled out by incorrect positioning of the reactive group. A recent example of a DNA-intercalating Zn(II) complex has been described by Yang and coworkers,42 who studied the reactivity of the Zn[(phen)(dione)Cl]ClO4 complex (Figure 13.15) toward pBR322 plasmid DNA. In this case the ligand itself is able to intercalate DNA with a binding constant of 2.4 × 104 M−1 and the cleavage proceeds with a rate constant of 5.8 × 10−5 s−1 at pH 8.1, 37 °C and a 3.0 mM complex concentration. The observed rate constant is relatively high, but it has to be taken in account that it has been obtained at high complex concentration. Due to the nature of the ligand, a direct comparison with a nonbinding complex is not possible, but the authors report that other complexes made by Zn2+ and phenanthroline are not active.

Conjugation with DNA-affine Subunits 383

Figure 13.15 The [Zn(phen)(dione)]2+ complex

+

+N

N N N

+N

Figure 13.16

N

2+

Zn

OH

N N +

Proposed structure of the active form of the Zn(II) complex studied by Mao

DNA affinity elements other than intercalators have also been used. Positively charged ammonium or peralkylammonium groups have a good affinity toward the polyanionic DNA. Schneider and coworkers43 studied the reactivity of the Co(III) complexes of cyclen derivatives bearing side chains of different lengths and terminating with a trimethyl ammonium group. At 37 °C and pH 7.0, they observed saturation kinetics, both with the Co(III)-cyclen complex and with the complexes of the alkylammonium conjugates. Interestingly, while the maximum reactivity remains constant within the series (kcat ca. 2 × 10−4 s−1), the binding constant increases with the length of the spacer (from 1.0 × 103 M−1 in the case of Co(III)-cyclen to 5.5 × 103 M−1 for the complex with a C6 spacer). Further increments of activity could be obtained by adding a second alkylammonium or Co(III)-cyclen groups. Again, peralkylammium groups were employed as affinity elements by Mao and coworkers.44 They synthesized two Zn(II) complexes of bipyridyl ligands bearing ethylammonium or butylammonium groups in the positions 5 and 5′. The former Zn(II) complex cleaves DNA at pH 7.2, 37 °C and 50 mM complex concentration with a rate constant of 3.8 × 10−5 sec−1. The pH reactivity profile shows a maximum at pH 7.5. On the basis of the crystal structure of the complex, the authors suggest that the quaternary ammonium groups (Figure 13.16) are placed in the right position to interact with the two negatively charged phosphate groups neighbouring the one that is activated by the metal ion. More sophisticated and biologically minded systems have been reported by the groups of Franklin and Sugiura. The first designed a 33-mer peptide incorporating a DNA recognition sequence and a metal-ion binding site.45 The peptide folds in a

384

Artificial Restriction Agents

helix-turn-helix (HTH) motif where the two helices have been derived from the DNA-binding engrailed homeodomain and the turn reproduces the sequence of the calcium binding site of calmodulin. Addition of Ce(IV) or Eu(III) leads to the formation of a peptide/lanthanide ion complex which retains the HTH tertiary structure. The Ce(IV) complex (and to a minor extent also the Eu(III) complex) promotes the cleavage of plasmid and linear double-stranded DNA and the reactivity of the complex is similar to that of the free ion. However, while the uncomplexed metal ion cleaves randomly both the P-O3′ and P-O5′ bonds, the peptide complex is regioselective for P-O3′ bonds. Furthermore, modest sequence selectivity for T/C rich sequences was also observed. In a similar approach, Sugiura and Nomura have succeeded in the preparation of zinc-finger peptides with hydrolytic ability toward phosphate esters, by mutating some amino acid residues coordinated to the zinc ion.46a This mutant is based on the sequence of the second finger in the three-tandem zinc finger protein Sp1, a DNA binding protein that binds specifically to GC boxes. Tandem alignment of three zinc finger mutants in a way similar to natural proteins resulted in efficient and site selective cleavage of plasmid DNA pUC19GC, and of a 37 bp DNA duplex, both containing a GC box. In another example, two zinc-finger peptides were connected to a peptide linker able to bind lanthanide ions.46b In the presence of Ce(IV) ions, the system cleaves linear dsDNA with high sequence selectivity.

13.6 Conjugation with Sequence-Selective Elements A more exciting and fascinating application of the DNA hydrolytic catalysts is the obtainment of artificial restriction enzymes with higher or different sequence specificity than natural systems. However, such a goal requires highly specific sequence recognition while most of the systems so far described produce random cleavage of DNA or show only modest sequence selectivity. A much higher selectivity is required and, in principle, this can be obtained by metal complexes conjugated with DNA oligonucleotides (for antisense recognition or triple helix formation) or PNA fragments. At the present time, only a very few examples of such systems are known. Komiyama and coworkers47 first reported, in 1994, a conjugate in which an iminodiacetate metal-binding group was appended to a 19-mer DNA oligonucleotide. In the presence of Ce(IV) ions, the DNA– iminodiacetate conjugate efficiently cleaved single-stranded 40-residue DNA at the linkage between residues 30 and 31, according to the predicted selectivity, with scission of the P-O5′ bond. Noticeably, the cleavage of the single-stranded DNA by this conjugate is much more efficient than hydrolysis of the dinucleotide TpT by Ce(IV)iminodiacetate, suggesting that the DNA moiety also has a role in delivering the metal ion close to cleavable phosphate groups. Krämer and coworkers described more recently a family of PNA–metal chelating group conjugates whose Zr(IV) complexes selectively cleave single-stranded DNA oligonucleotides.48 The cleavage is less efficient than in the case of Komiya-

The ARCUT System 385

ma’s DNA conjugate, probably due to the use of the less reactive Zr(IV) ion. Moreover, the employment, under the conditions used, of a large amount of free Zr(IV), due to the low metal affinity of the ligand (TRIS) subunit, leads to a substantial nonspecific random cleavage. At any rate, the fragments formed are consistent with the anticipated selectivity and the scission occurrs with remarkable regiospecificity at the P-O3′ bond.

13.7 The ARCUT System Very recently, Komiyama and coworkers described a new approach to the selective hydrolytic cleavage of DNA, based on the selectivity of the Ce(IV)-EDTA complex.5 As mentioned before, this complex cleaves ssDNA with such preference that bulges or gap-sites in dsDNA are selectively cleaved.17,18 In the ARCUT protocol, gap-like structures are formed at the desired sites of dsDNA or plasmid DNA by invasion of two pseudo-complementary PNA strands (pcPNA), whose sequences are selected in order to bind laterally shifted base sequences in the target (Figure 13.17).49 The two single-stranded portions formed in the invaded DNA are hence cleaved by incubation with Ce(IV)-EDTA. The sequence selectivity can be programmed without limitations and cleavage does not occur even in the presence of a single base mismatch.50 Cleaving efficiency is greatly enhanced by attaching a serine phosphate monoester at the pcPNA termini close to the gap site, since the dianionic terminal phosphate induces the accumulation of the Ce(IV) complex.50 However the reaction still requires, in many cases, high temperature (50 °C) and prolonged reaction times (64 h). DNA fragments obtained can be religated and connected with foreign double-stranded DNA by using DNA ligase and a ligation joint to provide recombinant DNA (Figure 13.17).18,49,51 The method has been applied by Komiyama and coworkers to the site-selective scission of genomic DNA of Escherichia coli (4.6 × 106 bases),50 to the site-selective scission of enzymatically methylated DNA (which is resistant to restriction enzymes)50, to the preparation of recombinant DNA18,49,51 and even of fusion proteins.52

gap sites Ligation joint

pcPNAs Ce (IV)/EDTA

Figure 13.17 binant DNA

Ligase

Schematic representation of the ARCUT protocol for the preparation of recom-

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Artificial Restriction Agents

13.8 A Critical Survey and New Perspectives Twenty years have passed since the first report by Barton and coworkers of an artificial agent for the hydrolytic cleavage of DNA.4 Since then, the realization of such systems has attracted great interest and has led, with Komiyama’s ARCUT,5 to the first examples of DNA manipulation performed with synthetic agents. Still, several important goals remain unachieved and the recognition of the most promising guidelines is still not easy. The ideal features of an artificial hydrolytic agent are high activity, high DNA affinity and sequence selectivity. Other requirements may vary depending on the final application: for an artificial restriction agent, double-stranded cleavage at the P-O3′ site is a requirement for subsequent religation, while for an antisense drug the absence of cofactors, activity at very low concentrations and at physiological pH and temperature are surely important. For all the possible applications, the occurrence of an oxidative cleavage, either as main or concurrent pathway, is highly undesirable. The last point is of particular relevance with synthetic agents. Metal ions used by nature in nucleases and phosphatases, e.g. Ca(II), Mg(II), Fe(II), Fe(III) and Zn(II), do not have, with exception of the Fe(II)/Fe(III) couple, relevant redox properties. Chemists, however, can pick up the most suitable candidate over the whole periodic table. The ideal metal ion for the realization of a hydrolytic system should have a hard character (to bind phosphate oxygen atoms), high Lewis acidity (to polarize both the phosphate group and the nucleophile) and a fast rate of ligand exchange (to ensure catalytic turnover).3o Several metal ions not used by nature best fulfil such requisites, particularly trivalent lanthanide ions, Ce(IV) and Cu(II). In the case of Cu(II), however, oxidative cleavage pathways have been detected for most of the hydrolytically active Cu(II)-based systems so far reported, both in the presence and in the absence of reducing agents and, on some occasions, even of oxygen. The adventitious presence of reducing agents either in DNA samples obtained from natural sources or in the in vivo environment, makes the use of Cu(II)-based systems as hydrolytic agents rather discouraging, notwithstanding their excellent activity. Moreover, the recent results form Rediijk and coworkers,9 who discovered oxidative DNA cleavage with a Zn(II) complex containing a phenol unit, put further emphasis on the need to assess clearly the reaction mechanism, even in the case of metal ions without a relevant redox activity. After the cleavage pathway, the first problem to face is that of the hydrolytic efficiency of the artificial system. A comparison of the activity of the different agents reported is rather difficult because of the large variety of reaction conditions used. However, a rough evaluation of the most reactive agents is tentatively shown in Table 13.1, which reports, where possible, the plasmid DNA degradation rate at a fixed agent concentration (5 mM, kψ′ ) and agent to DNA (base pair) ratio (0.125, kψ′′), and the Michaelis–Menten parameters. A first glance at the kψ′ and kψ′′ data reported, indicates that simple metal ions and monometallic complexes can hardly achieve relevant reactivity. Such low activity appears to be related mainly to a low affinity for DNA. Indeed, when the kcat

2.6·104 1.7·104 – – 1.3·104 3.0·103 1.0·103 – – – 5.5·103 – 2.5·105 1.7·103 – 1.0·104

7.0·10−5 1.0·10−4 – – 1.2·10−3 2.8·10−4 1.0·10−3 – – – 2.7·10−4 – 5.2·10−4 5.8·10−4 – –

Eu(III) Er(III) Zn(II)-pseudopeptide (50 °C) Zn-quercetin Cu(II)-TACH (35 °C) Pr(III)2-azacrown Er(III)2-shiff macrocycle Ce(IV)2-HXTA Fe(III)2-DTPB Zn(II)2-peptide Co(III)-cyclenC6ammonium Zn(II)-bipy-ammonium Cu(II)-neamine Eu(III)-naphtocrown Zn(II)-Rh-peptide Zn(II)-TACH-anthraquinone

8.0·10−6 7.8·10−6 ∼4·10−5 (3 m M) 1.7·10−4 (100 µ M) 7.3·10−5 4.1·10−6 5.0·10−6 1.4·10−4 (10 m M) 2.1·10−3 (100 m M) 1.0·10−5 (3.6 m M) 7.2·10−6 3.8·10−5 (50 m M) 2.8·10−4 4.9·10−6 2.5·10−5 (5.0 m M) 4.6·10−6 (5.0 m M)

kψ′ (s−1)b 4.1·10−6 3.8·10−6 – – 1.3·10−5 2.0·10−6 – 1.4·10−4 (r = 0.07) – 1.0·10−5 (r = 0.3) 3.5·10−6 – 3.1·10−4 2.3·10−6 2.5·10−5 1.9·10−6

kψ′′ (s−1)c

7.3 7.0 6.0 7.0

7.0 7.0 7.0 7.2 8.1 7.0 7.0 8.0 7.0 7.0 7.0

pH

c

b

19 19 12 40 7 19 – 150 77 12 19 38 50 19 40 12

[bp]d (mM)

Reactivity at 37 °C unless otherwise stated. Pseudo first-order rate constant for the cleavage of plasmid DNA at a fixed concentration of metal complex (5 µM). Pseudo-first-order rate constant for the cleavage of plasmid DNA at a fixed metal complex to DNA (base pair) ratio (0.125), unless otherwise stated. d Concentration of plasmid DNA reported as base pairs.

a

Ka (M−1)

kcat (s−1)

Hydrolytic Agenta

Table 13.1 Kinetic parameters for some of the most active artificial metallonucleases

39 11 20 21a 23 27 28 29 33 35 43 44 25 39 38 41

reference

A Critical Survey and New Perspectives 387

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values reported are examined, several agents appear have the potential to reach a good reactivity provided their affinity for DNA is increased. As inspired by natural enzymes, the development of bimetallic systems should be a promising strategy in order to increase the activity of the artificial hydrolytic agents. In these systems, not only the intrinsic reactivity may increase as a result of the cooperative action of the two metal centres, but also the DNA affinity should increase due to the larger positive charge of the complex and to the possibility of multipoint interaction with the DNA phosphate backbone. As a matter of fact, two of the most reactive agents so far reported, Liu’s Fe(III)2-DTPB33 and Que’s Ce(IV)2HXTA,29 are bimetallic. This strategy is particularly important, both in the case of Zn(II)-based systems, as the intrinsic hydrolytic activity of this metal is somewhat lower that that of the other ions, and in the case of lanthanide systems, since the obtainment of nonlabile metal complexes with free coordination sites usually leads to the use of polycarboxylic ligands which, as a drawback, reduce the reactivity of these ions. Nevertheless, the binuclear agents so far reported are relatively few and, in many cases, the reactivity gains are not so impressive when compared to the corresponding mononuclear complexes. Studies on model phosphate esters clearly indicate that the design of such systems is more delicate than the simple synthesis of ligands capable of binding two metal ions. The intermetallic distance and the rigidity of the complex are crucial features in order to allow full cooperation between the metal centres. Moreover, at least in the case of Cu(II)- and Zn(II)based agents, the formation of m-hydroxo bridges between the two ions can dramatically decrease the reactivity and must, accordingly, be strictly avoided.24a,53 Besides the realization of bimetallic systems, the main route toward the obtainment of efficient artificial agents requires, as mentioned before, an increase in the affinity of the hydrolytic complexes for DNA. The mechanism of the metal-ioncatalysed hydrolysis of phosphate esters involves, as a crucial initial step, the coordination of the substrate to the metal ion. This coordination is an essential requisite in order to deliver the metal-ion-coordinated hydroxide nucleophile close to the phosphate group, thus offsetting the electrostatic repulsion between the two negatively charged species, and allowing the activation of the phosphate toward nucleophilic attack. The metal-ion binding ability of phosphate diesters is rather poor and it is only slightly favoured by the polyanionic nature of DNA. It is not accidental that all the most reactive systems so far reported feature DNA affinity elements, such as two metal ions, intercalators, positively charged ammonium groups and aminosugars. Even in the cases where DNA binding has not been investigated, such as the Zn-(quercetin)2 complex,21 an effect of the poliphenolic ligand to increase the substrate affinity is probably present. However, the results reported indicate that the simple conjugation of a metal complex to a DNA-binding unit is not sufficient, particularly in the case of intercalating groups, and that a very careful design of the catalyst structure is required. In fact, in order to obtain the desired activity, the binding must lead to the correct geometry favouring a close contact between the metal complex and the DNA phosphate backbone. In this context, it appears that the design of the linker that connects the metal ion complex to the DNA-binding subunit is of great importance and, in particular, its length and flexibility. This is not an easy task since these features may easily change depending on the DNA binding site (major groove, minor groove, mismatches) and need to be tailored accordingly. In

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contrast, the results obtained by Ichikawa20 and Mao44 with Zn(II) complexes and by Schneider43 with Co(III) complexes bearing ammonium groups indicate that the use of such affinity elements may be much less demanding in terms of structural requirements. Short DNA oligonucleotides (for both antisense recognition and triple helix formation) or PNA fragments are probably the most promising DNA-binding subunits available to chemists. Indeed, such systems conjugate tight binding to DNA with high selectivity, and such features are ideally suited for the obtainment of a truly useful artificial nuclease. Such systems have been extensively studied for RNA3b but, up to today, very few agents have been studied with DNA and they have only been tested on short single strand fragments, while the problem of double-strand cleavage of DNA has still to be addressed.

13.9 Conclusions Since the first report on the ability of metal ions to promote DNA hydrolysis, tremendous progress toward the obtainment of efficient synthetic DNA hydrolytic agents has been attained. At present, chemists have at their disposal catalysts that cleave DNA in a few tens of minutes and, in some cases, can display interesting selectivity. However, several other aspects need to be addressed before a catalyst may reach the stage of practical application. Among the most important are: selectivity, double-strand scission and cleavage efficiency. The last point is of particular importance in the case of intrinsically less reactive, but biologically relevant metal ions, such as, for example, Zn(II). The use of multinuclear complexes as catalysts and of selective DNA-binding subunits supported by careful design of the system appear to be the main routes that have been successfully followed to date. Now, the problem is the integration of these different features into a single system in order to obtain really effective metallonuclease models. The Komiyama ARCUT system5 outlines an alternative strategy to that followed by the majority of the researchers in the field. In this case, selective cleavage of ssDNA over dsDNA has been exploited trough an extensive investigation of the reactivity of a single, very simple, and even moderately efficient cleaving agent. Such selectivity, conjugated with a smart design for the working protocol, has led to the realization of the only artificial restriction agent so far reported. Also in this case higher reactivity would be desirable in order to achieve shorter reaction times and higher cleavage yields, but the success obtained represents strong encouragement toward a renewed research effort in this field.

Acknowledgements We are deeply indebted to Prof. Umberto Tonellato, Prof. Paolo Scrimin, Prof. Manlio Palumbo, Dr Claudia Sissi and Prof. Stefano Moro for their productive collaboration and numerous stimulating discussions.

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14 New Metallo-DNAzymes: Fundamental Studies of Metal–DNA Interactions and Metal Sensing Applications Zehui Cao and Yi Lu

14.1 Introduction DNA was long considered only a carrier of genetic information. In 1994, Breaker and Joyce discovered that DNAs isolated using a combinatorial chemistry approach called in vitro selection,1–5 were also able to carry out catalytic activities.1 Since then, many catalytic DNA molecules, termed deoxyribozymes or DNAzymes, have been reported. Being the latest member of the enzyme family after protein and RNA enzymes, DNAzymes have rapidly demonstrated their capability of catalysing a variety of reactions previously known to only ribozymes or protein enzymes,6–9 including RNA/DNA cleavage,1,10–16 ligation,17–19 phosphorylation,18,20 cleavage of phosphoramidate bonds21 and porphyrin metallation.22 Figure 14.1 shows a few examples of DNAzymes. In addition, DNAzymes have shown catalytic efficiencies comparable to those of ribozymes or protein enzymes. For example, the ‘10-23’ DNAzyme identified by Santoro and Joyce has an observed catalytic efficiency of 109 M−1 min−1.15 Unlike RNA and protein enzymes, DNAzymes have not been found in nature, and in vitro selection remains the primary way to isolate and identify new DNAzymes. Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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3' N N N N N N Y rR N N N N N N N 5' 5' N N N N N N R N N N N N N N 3' A G G G C C T A A A G C CTA

3' N N N N N N G rN N N N N N N N 5' N N N N N N N 3' 5' N N N N N N T AA C C A G GG T G C G C C

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Figure 14.1 DNAzymes for Mg(II) (top), Pb(II) (middle) and Cu(II) (bottom). In each figure, ‘N’ represents any nucleotide provided they abide by Watson–Crick base pairing. All the DNAzymes contain an enzyme strand (bottom strand) and a substrate strand (top strand). Cleavage and ligation sites are marked by an arrow. Y denotes pyrimidine and R denotes purine. (Yi Lu, Prof; New Transition-Metal-Dependent DNAzymes as Efficient Endonucleases and as Selective Metal Biosensors. Chemistry – A European Journal, 2002, 8, 4589–4596. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Reproduced with permission.)

14.2 Metal Ions as Important Cofactors of DNAzymes Protein-based enzymes are constructed from 20 amino acids, whereas the variety of structural features of nucleic acid enzymes is limited to only four different nucleotides. The lack of the 2′-OH functional group in DNA compared to RNA further reduces the diversity of conformations that DNAzymes can take for effective target recognition. Nature’s solution to low efficiency and diversity in biomolecular interactions is the employment of cofactors. For example, protein enzymes are known to recruit cofactors, such as NADH, porphyrins and especially metal ions, to broaden the scope of reactions catalysed, increase catalytic efficiency and fine-tune reactivity.23–25 Therefore, to compete favourably with other families of enzymes, cofactors are even more critical for the functions of DNAzymes. Among common cofactors, metal ions are arguably the most important because of their capability to broaden and fine-tune the activity of the emzymes.23–25 In fact, other than a few reports that some DNA/RNAzymes are active independent of the presence of metal ions,13,14 for the majority of DNA/RNAzymes, metal ions, including Mg2+, Mn2+ and Ca2+, are

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essential for catalytic function at physiological conditions. However, most DNA/ RNAzymes found to date cannot match protein enzymes in terms of either the diversity of metal ions they employ or the specificity of the metal-binding sites. For example, protein enzymes use a wide variety of metal ions, including even the second and third-row transition-metals such as Mo and W, while metal ions used by DNA/RNAzymes are often limited to Mg2+, Ca2+ and Mn2+. Furthermore, it is well known that many protein enzymes possess remarkable metal-binding affinity and specificity, and they are commonly classified by the metal ions they specifically bind (e.g., copper proteins or zinc proteins). DNA/RNAzymes have also been shown to bind certain metal ions selectively, although the number of different metal ions that DNA/RNAzymes can bind is much fewer and the corresponding metal binding affinity is generally lower than those in metalloproteins.26–32 These weaknesses in DNA/RNAzymes are being overcome by using in vitro selection (see below).

14.3 Selection of DNAzymes Using in vitro Evolution Since DNAzymes are almost exclusively generated via in vitro selection, the key to more diverse metal ion cofactors, as well as higher binding affinities and selectivities is improving the selection process. Illustrated in Figure 14.2, generally the selection starts with a pool containing 1014 to 1015 random DNA sequences. Besides a random region in the middle, every DNA in the pool also contains two common priming regions at two ends for polymerase chain reaction (PCR). Under given conditions and cofactors, the DNA sequences that can carry out desired catalytic functions are separated from inactive ones. These active sequences are then amplified by PCR for the next round of reaction and selection. The stringency of the selection can be

Figure 14.2 (Plate 12) Schematic representation of the in vitro selection process (See colour plate section)

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enhanced in each round by reducing reaction time and cofactor concentration, so that highly active DNAs can be enriched gradually. This process is repeated for a certain number of cycles until DNA sequences with very high activity and selectivity can be isolated and identified. The same strategy has also been successfully employed in the field of aptamer research, where DNAs with high binding affinities and selectivities for the targets of interest, instead of high catalytic activities, are produced. The selection process has allowed the discovery of aptamers for a wide variety of targets, ranging from metal ions33 and small organic molecules,34,35 to proteins36 and even cancer cells.37,38 Theoretically, the selection of DNA/RNAzymes can also be carried out in the presence of many different cofactors, thus allowing the isolation of DNA/RNAzymes with very specific requirements for various cofactors, including any metal ions of interest. Indeed, efforts in this direction have yielded DNA/ RNAzymes that are catalytically active in the presence of some less common metal 43 ion cofactors, such as Pb2+,1,39 Cu2+,12,17 Zn2+,40,41 Co2+,42 and UO2+ It is expected 2 . that DNA/RNAzymes dependent on other metal ions can be generated by a similar approach with well-controlled selection conditions. An inherent problem with the in vitro selection strategy described above is that even though aptamers or DNA/RNAzymes can be obtained under controlled conditions, it does not exclude the possibility that these selected products can also function under different conditions or with different cofactors. This problem obviously results in lower-than-ideal selectivity of the aptamers or DNA/RNAzymes. In some cases, the activity of the nucleic acid enzymes in the presence of the target metal ion is even lower than with certain other metal ions. For example, the Zn(II)-binidng aptamer binds some other metal ions equally well, including Ni(II) and Co(II).44–46 Another DNAzyme, termed ‘10-23’ DNAzyme was selected with Mg(II) present, but its activity with Mn(II) is even higher.15,47,48 Furthermore, the same ‘8-17’ motif of a DNAzyme was obtained by four different laboratories with different cofactor conditions.15,49,40,50 It has a metal-dependent activity in the order Zn(II) >> Ca(II) > Mg(II) under similar conditions. Interestingly, further evaluation of this enzyme showed even higher activity with Pb(II).39 The apparent Kd values for Pb(II), Zn(II) and Mg(II) are 13.5 mM (at pH 6.0), 0.97 mM (at pH 6.0) and 10.5 mM (at pH 7.0), respectively.39 The lack of specificity in these DNAzymes poses major problems in studying metal DNA/RNA interactions and in applying DNAzymes as metal ion sensors. To solve this problem and improve metal ion specificity, a negative selection strategy can be introduced into the selection process by incubating the DNA pool with nontarget metal ions following positive selection with the target ion. In this step, any DNA sequences that have activity in the presence of other metal ions will be discarded, leaving the sequences that only function with the target ion present. This negative selection can be conducted multiple times as needed to greatly eliminate nonspecific sequences. As a demonstration of improved selectivity by negative selection, two parallel selections of Co(II)-dependent DNAzymes were carried out with and without negative selections (shown in Figure 14.3).42 In the absence of the negative selection, the resulting DNAzymes were more active with Pb(II) and Zn(II) than with Co(II). In comparison, when negative selection was incorporated, the obtained DNAzymes were much more active with Co(II) than with other metal ions. No detectable cleavage activity was observed with several other metal ions

Nucleic Acid Enzyme–Metal Ion Interactions 399 Initial pool of DNA/RNA Selections using only Co2+ DNA/RNA with Co2+ activity Positive Selection Selection using only Co2+

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Figure 14.3 Schemes of parallel selections of Co(II)-dependent DNAzymes with and without negative selection. (Yi Lu, Prof; New Transition-Metal-Dependent DNAzymes as Efficient Endonucleases and as Selective Metal Biosensors. Chemistry – A European Journal, 2002, 8, 4589–4596. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Reproduced with permission.)

investigated including Ca(II), Mg(II) and Cd(II), and cleavage activity in the presence of Mn(II) was minimum. This result shows the effectiveness of negative selection in improving specificity of DNAzymes and further demonstrates that in vitro selection is a powerful and versatile method for generating DNA/RNAzymes with the desired properties by carefully designed strategies and well-controlled conditions.

14.4 Understanding Nucleic Acid Enzyme–Metal Ion Interactions Compared to ribozymes and protein enzymes, the understanding of the detailed interaction between metal ions and DNAzymes and how metal ions affect the activities of the DNAzymes remains very limited. For instance, while many threedimensional structures of ribozymes and protein enzymes are available,51–54 only one DNAzyme structure has been reported.55 Yet this structure is not the enzyme’s active conformation, and thus provides limited insight into the enzymatic activity. Furthermore, whereas a great deal of knowledge has been available on the roles of metal ions in protein enzymes, much less is known about that in nucleic acids. As a result, understanding of the structure and function of metal ions in DNA/RNAzymes

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can bridge the knowledge gap between protein and nucleic acid enzymes, and also provide critical information for metal sensor development in sensing applications. Many techniques can be used to probe nucleic acid–metal ion interactions. Most of the approaches were initially designed and applied to study functions and mechanisms of ribozymes, because these catalytic RNAs are believed to play an important role in the early stage of life on Earth. One of the most intriguing objectives of nucleic acid enzyme research is the elucidation of structures of the active enzymes bound to metal ion cofactors. This goal was achieved by means of Xray crystallography. Crystal structures of metal ion–ribozyme complexes were obtained.56,57 These structures show that catalytic RNAs have complex three-dimensional structures and may contain multiple metal ion binding sites required for proper folding and activity. Even though X-ray crystallography provides direct visualization of nucleic acid enzyme structures, its application is limited to the solid crystal phase and often affected by the lack of diffraction quality crystals. Several other techniques, including nuclear magnetic resonance (NMR) and electron paramagnetic resonance (EPR) can be employed to study enzyme–metal ion interactions in the solution phase in close to physiological conditions. They often take advantage of paramagnetic metal ions or atoms on the enzyme, and monitor the changes in their spin states in magnetic fields during metal–enzyme interactions. For example, DeRose and colleagues used paramagnetic Mn2+ to substitute native Mg2+ in a hammerhead model ribozyme.58 The EPR signal from Mn2+ was sensitive to the environment surrounding the metal ion, thus providing small but measurable changes in the EPR signal during interaction with the enzyme. Specifically, binding of Mn2+ to a biomolecule induced a small ligand field asymmetry and increased its rotational correlation time, leading to considerable reduction in the amplitude of the six-line EPR signal from the bound Mn2+. This correlation was utilized to differentiate free and bound Mn2+ in the presence of the ribozyme. Binding affinity and the number of bound Mn2+ ions were determined using titration curves based on EPR signals. They found that the ribozyme contained four high-affinity binding sites for Mn2+ with a Kd of ∼4 mM and five other binding sites with a Kd of ∼460 mM. In a more recent report, Britt, Derose and coworkers employed pulsed EPR techniques of electron spin-echo envelope modulation (ESEEM) and electron spin-echo electron nuclear double resonance (ESE-ENDOR) for structural analysis of the paramagnetic metal ion Mn(II) bound to nucleotides and nucleic acids.59 Spin-echo EPR techniques allow the analysis of paramagnetic metal ion interactions with magnetic nuclei of ligands and nearby regions of bound macromolecules.59 A combined analysis of ESEEM and ESE-ENDOR data provided information on the number, type and spatial distribution of magnetic nuclei in the neighbourhood of the Mn2+ ion.59 Additionally, the exact metal ion ligation site on the nucleotide and the hydration number of Mn2+ in the complex were also determined. In another related report, Schiemann and coworkers employed a strategy that replaced Mn2+ ions bound to a Diels–Alder ribozyme with higher-affinity, but EPR-silent, Cd2+ ions. In this way, they were able to characterize each of the five Mn2+ binding sites individually using EPR signals.60 Given these results, it is clear that EPR and related techniques provide a powerful approach to highly precise characterization of the structural features of nucleic acid–metal ion complexes.

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Other methods have also been developed to study nucleic acid–metal ion interactions. Pyle and coworkers used terbium(III) to identify metal ion binding sites of a family of group II intron ribozymes.61 Tb3+ and related ions are able to cleave RNA efficiently under physiological conditions. In this study, various concentrations of Tb3+ were incubated with the ribozymes in the presence of Mg2+. The cleaved fragments of the enzymes were analysed by polyacrylamide gel electrophoresis (PAGE). Fragments at low Tb3+ concentrations revealed strong metal ion binding sites, while fragments produced by high concentrations of Tb3+ corresponded to lower-affinity binding sites. Comprehensive analysis of the gel data was successfully used to map most metal ion binding sites of the ribozymes. These Tb3+-binding sites were found to colocalize with Mg2+-binding sites based on two results. First, addition of Mg2+ at low pH effectively eliminated Tb3+ cleavage. Second, Mg2+ was shown to cleave at the same Tb3+ cleavage sites, only at much slower rates. In addition to mapping metal ion binding sites by metal-ion-mediated hydrolysis, NMR and related techniques have been useful in studying nucleic acid enzyme– metal ion interactions. Sigel, Pyle and coworkers employed NMR and two-dimensional nuclear overhauser enhancement spectroscopy (NOESY) to reveal the solution structure of the central core of a group II intron ribozyme and its interactions with Mg2+.62 It was found that binding to Mg2+ had most impact on the nucleotides in the bulge region of Domain 5. Using NMR and other techniques, Sigel and coworkers reported the solution structure of a catalytically active Domain 6 (D6) of a selfsplicing Group II intron ribozyme. A single unpaired adenosine was found to reside within the helix of D6 and was partially stacked between two flanking GU wobble pairs. A novel prominent Mg2+ binding site was identified in the major groove of this site. In a recent report, Sigel and coworkers performed paramagnetic linebroadening experiments with Mn2+ and titration studies with Mg2+ using NMR to determine the affinity constants for Mg2+ binding to five distinct sites in Domain 6 of Group II introns.63 Even though most of the aforementioned methods have been developed and used for ribozymes and RNAs, adaptation of these techniques to DNAzymes is not likely to cause major difficulties. Many efforts are underway and possibilities are being explored along this line of research. Compared to many other techniques, methods based on fluorescence resonance energy transfer (FRET) have been shown to be able to provide unique information on structural alterations of the DNA/RNAzymes upon metal ion binding in real time with minimum perturbation of the interactions. FRET is the energy transfer between two fluorophores when they have inherent spectrum overlap and are in close proximity (<10 nm). The result is that when excited, the donor fluorophore generates less fluorescence output compared to when there is no FRET, while fluorescence emission from the acceptor fluorophore is observed even though it is not usually excited by the donor excitation. Since the FRET efficiency is tightly related to the distance between the two fluorophores, it can often be used to report subtle intramolecular distance changes or structure alterations. To study DNAzyme–metal ion interactions in real time, an initial FRET study was carried out with a trifluorophore-labelled DNAzyme,64 followed by a more comprehensive study of the metalion-induced folding of DNAzymes using FRET (Figure 14.4).65 In the latter report,

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Figure 14.4 FRET efficiency and catalytic activity (kobs) of the 8-17 DNAzyme in the presence of Zn(II) (A), Mg(II) (B) and Pb(II) (C), and proposed folding model (D). (Reprinted with permission from J. Am. Chem. Soc., 2007, 129, 6896–6902. Copyright 2007, American Chemical Society.)

the Cy3/Cy5 FRET pair was labelled each time on two of the three stems of the Pb2+-dependent ‘8-17’ DNAzyme. The FRET efficiency between Cy3 and Cy5 was monitored as an indication of the distance change between the two stems when metal ions were introduced into the DNAzyme solution. FRET results indicated that, in the presence of Zn2+ and Mg2+, the DNAzyme folds into a compact structure, in which Stem III approached a configuration defined by Stems I and II, without changing the angle between Stems I and II. Correlations between metal-induced folding and activity were also studied. For Zn2+ and Mg2+, the metal ion with higher affinity for the DNAzyme in global folding (Kd(Zn) = 52.6 mM and Kd(Mg) = 1.36 mM) also displayed higher affinity in activity (Kd(Zn) = 1.15 mM and Kd(Mg) = 53 mM) under the same conditions. Global folding was saturated at much lower concentrations of Zn2+ and Mg2+ than the cleavage activities, indicating the global folding of the DNAzyme occurs before the cleavage activity of the metal ions. Surprisingly, no Pb2+-dependent global folding was observed. These results suggest that for Pb2+, global folding of the DNAzyme may not be a necessary step in its function, which may contribute to the DNAzyme having the highest activity in the presence of Pb2+. While FRET studies in bulk solutions offer useful insights into DNAzyme folding and activity, some valuable information may be lost due to the averaging effect of the large quantity of molecules present in the solution. Moreover, higher temporal resolution is needed in order to reveal the real underlying mechanism of the metal-ion-dependent catalytic activities. To overcome these limitations, single-

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Figure 14.5 (Plate 13) Single molecule FRET study of the 8-17 DNAzyme: (a) time traces of the FRET signals and changes upon injection of 100 mM Zn2+ at 21 s; (b) distribution of single DNAzyme molecules with different FRET efficiencies before and after injection of 20 mM Pb2+; (c) proposed reaction pathways of the DNAzyme in the presence of Zn2+/Mg2+ and Pb2+. (Reprinted with permission from Nat. Chem. Biol., 2007, 3, 763–768.) (See colour plate section)

molecule FRET (smFRET) was developed and applied to the study of the ‘8-17’ DNAzyme (Figure 14.5).66 Divalent metal-dependent folding, cleavage and release of the product of the DNAzyme probed at the single molecule level, combined with statistical analyses, provided a unique opportunity to dissect the reaction pathways and to determine the rates of both cleavage reaction and product release unambiguously. This study reveals that the catalytically active DNAzyme does not undergo global folding before cleavage if Pb2+ is used as the cofactor. Upon addition of Zn2+ and Mg2+, the reaction proceeds through a folding step followed by a cleavage reaction, suggesting that the DNAzyme may require metal-dependent global folding for activation (Figure 14.5). The different reaction pathways between Pb2+-dependent and Zn2+- and Mg2+-dependent reactions may contribute to the high activity of the Pb2+-dependent 8-17 DNAzyme, which is among the fastest nucleic acid enzymes. These results suggest that besides the induced-fit mode of activation found in many DNAzymes, the ‘8-17’ DNAzyme may be a rare case where the much faster lockand-key mode is also possible.

14.5 DNAzymes as Metal Ion Sensors Metal ions often have profound impact on human health. Their effects on health are usually concentration dependent. While many metal ions are essential for

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functions of human body at low levels, over-exposure to these ions can be detrimental. For example, iron is a critical component of the transportation system for oxygen and electrons in the human body, and is tightly associated with many enzymes. Deficiency in iron leads to anaemia, which is especially harmful to pregnant women.67 On the other hand, it has also been shown that excess ‘free iron’ in the human body can produce oxidative stress, and is involved in many pathological manifestations (arteriosclerosis, diabetes, aging, etc.).68 Therefore, it is clearly beneficial to be able to monitor levels of different metal ions as part of disease diagnoses or treatment plans. In a related issue, since metal ions can enter the human body through contact with the environment, for example drinking water, it is essential to monitor and control metal ion levels in these sources to reduce metal-related health risks. Laboratory techniques, such as atomic absorption spectrometry, inductively coupled plasma mass spectrometry (ICP-MS), anodic stripping voltammetry, kinetic phosphorescence analysis, X-ray fluorescence spectrometry and microprobes, have been routinely used for metal ion analysis with high sensitivity. Many of them can simultaneously quantify multiple metal ions. However, most of the above techniques require sophisticated equipment and sample pretreatments, or skilled operators, making it difficult and quite costly for on-site, real-time monitoring. To circumvent these problems, DNAzymes have been developed as metal sensors because of their metal-ion-dependent catalytic activities. DNAzymes have many advantages as metal ion sensors compared to many conventional metal detection techniques. First, DNAzyme sensors can reduce the requirement for complex and expensive instruments, and can be built into a portable package for on-site monitoring. Second, DNAzymes are selective, due to the dependence on specific metal ion cofactors. Their catalytic activities can be readily engineered to generate and amplify detectable signals for sensitive analysis. DNA-based sensors are also environment-friendly by being biocompatible and biodegradal, and should pose minimum toxic effects on the human body. Compared to antibody- and RNA-based sensors, DNAzymes are more resistant to hydrolysis and stable for long-term storage and application. They can even be denatured and renatured for many cycles without losing much of the activitiy. Lastly and perhaps most importantly, DNAzymes provide a common platform for developing sensors for new metal ions of interest or even targets beyond metals. Once the strategy of signal transduction to convert the catalytic activity to measurable signals is developed, the same design can be readily applied to another target as soon as a DNAzyme is selected for the target. This flexible platform is expected to reduce the cost of developing sensors for different targets in large quantities, and make these sensors more suitable for parallel and multiple detection of complex samples in an array format.

14.5.1 Fluorescence-Based DNAzyme Sensors Based on Catalytic Beacons A fluorescence-based lead sensor was initially constructed based on the ‘8-17’ DNAzyme (Figure 14.6).39 Fluorescence was employed because of its proven high sensitivity. An enzyme DNA strand and a substrate strand were hybridized to form

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Figure 14.6 A catalytic beacon Pb2+ sensor: (Left) Schematics of beacon signal generation; (Right) selectivity of the sensor, inset: kinetics of fluorescence increase with Pb2+ or other divalent metal ions. (Reprinted with permission from J. Am. Chem. Soc., 2000, 122, 10466– 10467.)

the lead sensor. Furthermore, a fluorophore was used to label the 3′-end of the enzyme strand and a fluorescence quencher to label the 5′-end of the substrate. Close proximity between the fluorophore and quencher resulted in a low fluorescence background. When lead was present, the enzyme was activated to cleave the substrate at the cleavage site. The smaller substrate fragment could not stay hybridized to the enzyme strand. Therefore these two strands were detached, causing separation of the fluorophore and quencher and consequently restored fluorescence. This sensor turned out to be highly sensitive for lead ions with a detection limit of 10 nM. It also responded >80 times better to Pb(II) than to the next competitive metal ion (Co(II) and Zn(II)) and hundreds times better to other metal ions tested. One problem with this sensor was that it required 4 °C for reasonably low fluorescence background and effective detection. To overcome this limitation, a new sensor design was proposed, which added an additional quencher at the 5′-end of the substrate strand.69 This internal quencher ensured a low background in the absence of lead even when the substrate was not efficiently hybridized to the enzyme. Indeed, a much improved signal-to-background ratio was obtained and sensitive lead detection could be conducted at room temperature. Two additional advantages of the fluorescent DNAzyme sensor are worth mentioning. First, since the detection is based on the kinetics of fluorescence enhancement, it is less affected by background fluorescence in environmental samples. Additionally, because catalytic reactions are responsible for signal generation, a single Pb2+ ion can catalyse the turnover of multiple fluorescent DNAzymes when even higher sensitivity is desired. This successful sensing strategy has been applied to other DNAzymes and their metal ion cofactors. One notable example is the sensor for UO2+ 2 recently developed in our lab (Figure 14.7).43 A DNAzyme specific for UO2+ was first identified via in 2 vitro selection. Following that, the same one-fluorophore-two-quencher approach was adapted to the new DNAzyme and UO2+ 2 detection was achieved with excellent sensitivity and selectivity. Most sensors based on molecular probes are not expected

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Figure 14.7 (Plate 14) Design and performance of DNAzyme-based UO22+ sensor. Competing metals were tested from 10 mM to 1 mM, and UO22+ (the last three bars) was tested from 0 to 10 nM. Inset: Sequence, schematics and fluorescent detection of the DNAzyme-based UO22+ sensor. (Juewen Liu et al., A catalytic beacon sensor for uranium with parts-per-trillion sensitivity and millionfold selectivity, Proc. Natl. Acad. Sci. USA, 104, 2056–2061. Copyright 2007, National Academy of Sciences, USA.) (See colour plate section)

to compete or exceed instrumental analysis. However, the UO2+ 2 DNAzyme sensor was able to realize a detection limit of about 45 pM, even lower than that obtained on ICP-MS (420 pM). In addition, it has a selectivity of more than one million-fold over any other metal ions tested. Development of fluorescent sensors for paramagnetic metal ions (i.e., those metal ions with unpaired electrons) such as Cu2+ has long been a challenge, due to their intrinsic fluorescence quenching properties that often cause decreased emission upon metal binding. On the other hand, DNAzyme-based sensors are less restricted by this problem since the metal-recognition core of the enzyme can be separated from the fluorescent signalling moiety, thus minimizing the contactinduced quenching. Therefore our laboratory developed a fluorescent DNAzyme sensor for Cu(II) based on a previously reported Cu(II) DNAzyme by Breaker and coworkers.12,70,71 This DNA-cleaving DNAzyme was tagged with a fluorophore and two quenchers similar to the sensors above.72 With this sensor, Cu2+ was readily detected down to 35 nM at room temperature, and the selectivity was 2000-fold better than other tested ions. We have recently designed a modified DNAzyme sensor for Hg2+ detection, using the dual quencher method to construct a catalytic beacon. Five T-T mismatches replaced the double-strand in the loop region of the UO2+ 2 DNAzyme that is required for binding UO2+ , thus inactivating the enzyme. Addition of Hg2+ could 2 73–75 stabilize the T-T mismatches, resulting in restored activity. This sensor has a detection limit of 2.4 nM, the lowest among known Hg2+ sensors based on small or macromolecules. The examples shown above all require labelling with fluorophores and quenchers on the termini of the DNA strands. However, the ideal positions to place the fluorophore and quencher pair would be on the two nucleotides closest to the cleavage site on the substrate strand. In this way, the quencher on the enzyme strand would no longer be necessary since the quencher and fluorophore on the substrate could stay close enough for minimum background. Cleavage of the substrate would then separate this pair and generate enhanced fluorescence. One concern for this design is possible activity decrease due to these two modifications near the active

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site. To investigate the feasibility of internal fluorophore/quencher labelling on DNAzyme, Li and colleagues conducted systematic studies by placing different dye/quencher pairs at various locations on the substrate.76 The results indicated that generally, the closer the dye/quencher to the cleavage site, the higher the fluorescence enhancement upon lead-induced enzymatic cleavage. However, the reaction rates were also lowered as a result. Given these observations, internal dye/quencher labelling for making DNAzyme sensors is feasible, but the design needs to be optimized. Instead of developing sensing strategies based on known DNAzymes, one appealing alternative would be directly selecting DNAzymes with a built-in signal transduction mechanism. Li and coworkers have realized this experimentally by modifying the general in vitro selection scheme.16,77 Compared to conventional selection, these authors added a few extra steps to each round to introduce a short nucleic acid strand to the DNAs in the pool before reaction in the presence of metal ions. This short strand contained an adenine ribonucleotide (rA) as the cleavage site, and a fluorophore and quencher labelling the two nucleotides closest to rA. When incubated with target metal ions, only those sequences that could cleave the rA site were collected and enriched. The final products of this selection were DNAzymes that were able to function normally with the internal labels and generate greatly enhanced fluorescence upon addition of target metal ions.

14.5.2 Colorimetric Sensors Based on DNAzymes Fluorescence-based DNAzyme sensors are excellent for metal ion detection due to their high sensitivity and selectivity. They also provide a solution to on-site metal analysis when equipped with portable fluorometers. However, to further reduce the cost and simplify the process of metal sensing, thus making it readily accessible to a much wider range of fields, including clinics, battlegrounds and even households, new strategies need to be employed. We have made progress in this direction with the development of colorimetric metal ion sensors. In a colorimetric sensor, a colour change is observed in the presence of target analytes. Compared to other techniques, colorimetric sensors can minimize or even eliminate the need for analytical instruments, making on-site detection much more convenient. Some common colour-reporting groups include small organic dyes, conjugated polymers and metallic nanoparticles. Among them, metallic nanoparticles display strong distance-dependent optical properties and very high extinction coefficients.78 Therefore, we designed our first-generation colorimetric lead sensor based on DNAzymes and DNA-functionalized gold nanoparticles (AuNPs).79 Briefly, the substrate of ‘8-17’ DNAzyme was extended at both ends with a short DNA strand of the same sequence. In this way, when AuNPs functionalized with the complementary DNA (cDNA) of the short extension were introduced, the AuNPs became literally crosslinked by the extended substrate to produce aggregated nanoparticle clusters (Figure 14.8). Such AuNP aggregates display a blue colour, due to their strong absorption of light with wavelengths around 700 nm. The ‘8-17’ DNAzyme was then introduced to hybridize with the remaining sequence of substrate. When Pb(II) was present, the enzymatic reaction was initiated to cleave the

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Figure 14.8 DNAzyme and AuNP-based colorimetric Pb2+ detection: (A) Pb2+-directed assembly of DNAzyme-linked AuNPs aligned in a head-to-tail manner; (B) colour of the AuNPs in the presence of different divalent metal ions. (Reprinted with permission from J. Am. Chem. Soc., 2003, 125, 6642–6643. Copyright 2003, American Chemical Society.)

substrate sequence, leaving a broken linkage between the AuNPs. Consequently, the dispersed AuNPs displayed a deep red colour due to a shifted absorption to around 522 nm. With increasing level of lead ions, a blue-to-red colour shift was clearly discernible by the naked eye. The detection of lead ions could also be realized more accurately by UV/Vis spectroscopy measurements. The ratio of extinction at 522 nm over 700 nm was used to quantify Pb(II) since it rose with increasing concentration of lead ions. A detection limit of ∼100 nM was achieved and the sensor was able to detect lead extracted from paint. An important benefit of the AuNP-based DNAzyme sensor is the easy implementation of tuneable dynamic ranges. A sensor design that allows simple tuning of dynamic range is highly desirable to match the requirements of different applications. For example, the US EPA defines the toxic threshold for lead in paint as ∼2 mM, while that in drinking water is 72 nM. Additionally, regulations on sensitivity of metal ion detection can change over time. It is often not economical or even practical to redesign and remake new sensors to meet these regulation changes, thus giving extra significance to tuneable sensors in practical applications. Taking advantage of the knowledge obtained from biochemical assays and the multiple turnover property of the DNAzyme, we have demonstrated a general scheme for tuning the dynamic range of the colorimetric Pb2+ sensor system.79 DNAzyme activity was abolished by changing the G•T wobble pair to a G-C Watson–Crick pair (Figure 14.9, left).80 This mutated DNAzyme could still assemble AuNPs similarly to the native DNAzyme. When a ratio of 5 : 95 was adopted for the active enzyme (17E) versus inactive enzyme (17Ec), the Pb2+-sensitive range shifted to ∼one order of magnitude higher Pb2+ concentrations (Figure 14.9, right). This tuning property is unique and useful because it allows detection of Pb2+ over a wide concentration range without redesigning the sensor.

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Figure 14.9 (Left) Secondary structure of the 8-17 DNAzyme for lead ion detection; (Right) dynamic ranges of the colorimetric sensors with different enzyme compositions. (Reprinted with permission from J. Am. Chem. Soc., 2003, 125, 6642–6643. Copyright 2003, American Chemical Society.)

In the first generation colorimetric sensor for lead, an additional heating and cooling step was needed before the DNA-functionalized AuNPs could be effectively crosslinked by the extended substrate strand. In fact, detection of lead ions was also conducted after this annealing step, where absence of lead produced a blue product and lead ions gave a red colour. This problem most likely arose from the alignment of the AuNPs. Based on the design, the substrate hybridized at both ends with two AuNPs tagged with the same cDNA sequence. This means these two AuNPs had to take a head-to-tail alignment, which induced a significant steric effect to prevent the crosslinking. To solve this problem, a tail-to-tail design was adopted and aggregation of AuNP was observed without the annealing step.81 Additionally, the size of the AuNP was changed from 13 nm to 42 nm, which greatly reduced the time needed to see a colour change to 5 min.81 Later, a true ‘turn-on’ colorimetric sensor was developed in which addition of lead ions could directly break up preaggregated AuNPs to produce a distinctive blue-to-red colour shift.82,83 To accelerate Pb2+induced disassembly, small pieces of DNA (called invasive DNA) complementary to the cleaved substrate strands were employed to facilitate AuNP release. Under optimized conditions, a blue-to-red colour shift was observed in 5 min at room temperature in the presence of Pb2+.82 Recently, we further optimized the disassembly conditions by designing asymmetric substrate binding arms so that disassembly occurred right after cleavage without the use of invasive DNAs.83,84 14.5.3 Other DNAzyme-Based Metal Ion Sensors The DNA backbone of the DNAzymes allows convenient modifications to the enzyme to tag a signalling moiety, facilitate immobilization of the enzyme or add additional functionality. Therefore, a variety of other detection strategies can be developed for metal ion sensing based on DNAzymes. In one such attempt, Plaxco and coworkers immobilized the lead-dependent 8-17 DNAzyme onto a gold electrode through the 5′-end of the enzyme strand and attached a methylene-blue (MB) group to the 3′-end.85 The DNAzyme remained rigid when hybridized with the substrate strand, separating the MB group from the surface and giving a low rate of electron transfer. In the presence of Pb2+, the substrate was cleaved and released,

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making the immobilized enzyme more flexible. Therefore, enhanced electron transfer was observed. Similar to other lead DNAzyme-based sensors, this electrochemical sensor was highly selective for Pb2+. A detection limit of 300 nM was reported and the selectivity was comparable to those of the fluorescent and colorimetric sensors based on the same DNAzyme. By immobilizing DNAzymes on a solid surface, an alternative sensing platform can be built. Notably, this scheme will allow sensor regeneration, as well as long-term storage of the sensor. In one report, the lead-specific 8-17 DNAzyme was covalently attached to gold surfaces.86 In the presence of Pb2+, a fluorophore-labelled substrate fragment was cleaved and released into solution for detection. Thanks to a very low background, a detection limit of 1 nM was achieved. In another report, surfaceimmobilized DNAzyme retained its activity even after storing in a dried state for 30 days at room temperature.87 A detailed characterization of this system was later carried out.88 These studies will facilitate the application of DNAzymes in sensor arrays for analysing complex targets.

14.6 Summary Studies of DNAzymes represent an actively developing interdisciplinary field involving many aspects of chemistry research, including biological, inorganic, physical and analytical chemistry. In some ways, this field has similarities to the study of ribozymes. However, it also holds many new possibilities in exploring structural mechanisms of functional nucleic acids and their biosensing applications. Despite recent progresses, there is still a clear lack of understanding of how DNAzymes carry out their catalytic functions, in comparison with our understanding of protein and RNA enzymes. The three-dimensional structure of the DNAzyme in its active form containing its cofactors has yet to be obtained. The structure features of the DNAzyme will greatly help elucidate the underlying mechanism of DNAzyme–cofactor interactions. In addition, analysis of the function of DNAzyme and its interactions with the surrounding microenvironment at the molecular level using EPR, NMR, smFRET, etc., is also in great need because they can provide valuable information toward the ultimate understanding of DNAzyme activities. On the other hand, given the current level of knowledge of DNAzymes, we have already been able to develop DNAzymes with the desired properties in the presence of specific cofactors using versatile in vitro selection. It will be of great significance to actively extend the family of DNAzymes in pursuit of a wider range of catalytic reactions that they can carry out and an increasing number of cofactors that they can recruit. Due to the fact that DNAs are inherently more stable than RNAs, these newly developed DNAzymes are more suitable to be engineered into practical biosensors for many different targets of interest. Additionally, with a large quantity of DNAzyme-based sensors available, comprehensive sensor arrays can be built for highly reliable analysis of complex samples. Given the increasing demand for convenient, economical yet effective sensors, we will continue to witness rapid development and advancment of the DNAzyme sensing field.

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Acknowledgements The Lu group research described in this review has been supported by the Department of Energy (DE-FG02-08ER64568) and the National Science Foundation (CTS-0120978, DMR-0117792, and DMI-0328162).

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15 Two-Metal-Ion-Dependent Catalysis in Nucleic Acid Enzymes Wei Yang

15.1 Chemistry of Nucleic Acid Synthesis, Cleavage and Strand Transfer DNA and RNA are synthesized by polymerases in the 5′ to 3′ direction. The hydroxyl group at the 3′ end of a primer strand attacks the a-phosphate of an incoming (deoxy)nucleoside triphosphate ((d)NTP) to incorporate the (d)NMP into the primer strand and release a pyrophosphate molecule (Figure 15.1A). Therefore, nucleic acid synthesis transfers a phosphodiester bond from a (d)NTP to the 3′-OH (the nucleophile) of a growing primer strand. All polymerases have been shown to require Mg2+ ions for catalysis.1,2 The same phosphoryl transfer reaction occurs during DNA or RNA cleavage except that the phosphate being attacked is the backbone of a nucleic acid and the nucleophile is a hydroxyl group from water, protein or ribose (Figure 15.1B,C). When a water molecule serves as a nucleophile, the nucleic acid is hydrolysed. When the nucleophile is a hydroxyl group of protein sidechains, e.g. Ser and Tyr, as in DNA recombination,3–5 a 2′-OH of ribose, as in RNA cleavage6 or a 3′-OH of RNA or DNA, as in splicing and integration,7,8 the phosphorylated product is covalently linked to the protein or nucleotide (Figure 15.1B). The chemistry of a phosphoryl transfer reaction begins with deprotonation and activation of a nucleophile and finishes with protonation of the leaving group. The mechanism for phosphoryl transfer reactions in nucleic acids has been shown to be the SN2 type, involving a pentacovalent phosphate intermediate (phosphate anion) and inversion of the stereo Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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Figure 15.1 Phosphoryl transfer reactions in nucleic acid synthesis, cleavage and recombination: (A) DNA and RNA synthesis. An incoming (d)NTP is incorporated into the primer strand. The 3′-OH of the primer strand is the nucleophile and attacks the a-phosphate of (d)NTP (highlighted in red). Pyrophosphate (PPi) is the leaving group. (Reproduced from Yang, W. et al., Making and breaking nucleic acids: two-Mg2+-ion catalysis and substrate specificity, Mol. Cell, 2006, 22, 5–13); (B) Cleavage. Scissile phosphate (highlighted in red) is transferred to water (hydrolysis), a nucleotide (as in splicing), a Tyr or Ser protein side chain (as in DNA recombination), or another DNA or RNA strand (as in RNA splicing or DNA recombination) depending on the nucleophile. The products are 3′-OH and 5′-PO4. (C) Same as (B) except the products are 5′-OH and 3′-PO4

configuration at the phosphorus.9–11 A nucleophile can attack the scissile phosphate from either side of the phosphate, and thus either the 3′- or 5′- OH can be the leaving group4,5,12 (Figure 15.1B,C). To date, except for catalysis that results in a 3′-phosphate and a 5′-OH,4,6 one or two metal ions are required for the phosphoryl transfer reaction. 15.1.1 Basic Properties of Mg2+ and Divalent Cations Mg2+ and Ca2+ are the most abundant divalent cations in living organisms,13,14 and Mg2+ is the most abundant divalent cation inside cells.15,16 Others, like Fe2+, and Zn2+ and Cu2+ are widespread, and Mn2+ and Ni2+ are essential, but found at low concentrations.14 The foremost quality of these divalent cations is the high density of positive charge, which makes them efficient for charge neutralization of phospholipids and nucleic acids. The second common property is hydration and the specific ligand requirement. By the atomic or covalent radius Ca2+ is the largest (1.9 Å), Mg2+ the second (1.6 Å) and the rest are similarly smaller (1.3 Å). However, when hydrated, Mg2+ is larger than Ca2+ because Mg2+ is associated with multiple shells of ligand water molecules.16 In biological systems these metal ions never exist without water

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Figure 15.2 Preferred coordination geometry of divavlent cations (Me2+): (A) Mg2+ prefers octahedral coordination with six ligands; (B) Zn2+ prefers tetrahedral configuration with four ligands. In protein structures, both Mg2+ and Zn2+ have been observed with five ligands in nonstandard geometry

or ligand. Therefore ionic (Shannon) radii17 are inapplicable. The most common ligand coordination geometry is octahedral or tetrahedral18 (Figure 15.2). Mg2+ prefers to have six inner-sphere ligands arranged in an octahedral configuration, and so does Fe2+.18 Ca2+ and Zn2+ can have octahedral coordination, but Zn2+ is frequently coordinated by four ligands in a tetrahedron,19 and Ca2+ by seven, eight or even nine ligands.20 The number and length of ligand bonds are empirically determined and shown to vary within a very narrow range.18 These metal ions are far more sensitive to the ligand type and geometry than hydrogen bonds or salt bridges between oxygen and nitrogen atoms. Each of these divalent cations has been found to be essential for catalysis by at least one enzyme, e.g. Fe2+ in nonheme iron enzymes (oxygenases and demethylases),21 Zn2+ in deacetylase and protease,22,23 Ni2+ in urease,24 but the Mg2+ ion is most frequently associated with nucleic acid enzymes.13 This is perhaps because of its abundance and solubility, stability in redox potentials compared with Mn2+, Fe2+ and Cu2+, its small size relative to Ca2+ and its rigid coordination geometry compared to the transition metals Fe2+, Cu2+, Ni2+ and Zn2+.16 The ligand bond of the Mg2+ ion is empirically determined to be 2.07 Å.18 The unusual hydration property of Mg2+ ions may also be a factor for its catalytic role. Mg2+ ions exhibit extremely slow exchange rates of inner-shell water molecules16,25 and often retain a couple of water ligands when coordinated by organic or macromolecules.18 The pKa value of a water molecule is reduced from 16.0 in free solution to 11.4 when associated with Mg2+.26

15.2 All DNA and RNA Polymerases Require Two Mg2+ Ions for Catalysis To date, six families of DNA polymerases have been identified, and each contains a number of conserved sequence motifs.27,28 The first four families, A to D,

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Figure 15.3 (Plate 15) Three types of catalytic centres found in DNA and RNA polymerases; (A) Phage T7 DNA polymerase represents the pol I-like active centre, which is found in the A, B and Y-family DNA polymerases, retroviral reverse transcriptases, phage RNA polymerases and viral RNA-dependent RNA polymerases. The catalytic residues (highlighted in blue and red) include two Asps and one Glu on two adjacent antiparallel strands (highlighted in cyan); (B) Human DNA pol b represents the pol b-like catalytic centre, which is found in the C and X-family DNA polymerases, poly-rA polymerases and CCA-adding enzymes. The catalytic residues include three conserved Asps on two adjacent parallel strands; (C) S. cerevisiae RNA pol II represents all bacterial and eukaryotic DNA-dependent RNA polymerases. The catalytic core consists of a b-barrel with three catalytically essential Asps on a loop (highlighted in cyan). All of these polymerases contain two absolutely conserved Asps, which together with the a-phosphate of (d)NTP jointly coordinate the two Mg2+ (shown as green spheres) (See colour plate section)

encompass all polymerases for DNA replication and a few for repair, while the remaining polymerases in the X and Y families are involved in DNA repair exclusively. Crystal structures of A, B, C, X and Y family polymerases have been determined.1,29–33 The structures of the catalytic domains and the active sites of these DNA polymerases fall into two classes, the pol I-like, which includes the A, B and Y family polymerases (Figure 15.3A), and the pol b-like, which includes the X and C family polymerases (Figure 15.3B). The catalytic domains of both classes contain a b-sheet and several a-helices, but the topology and locations of the catalytic residues differ. In the pol I-like polymerases, the central b-sheet is antiparallel and the catalytic residues are located on two adjacent strands, 1 and 3 (Figure 15.3A). In the pol b-like polymerases, the b-sheet is mixed parallel and antiparallel and the catalytic residues are located on two adjacent parallel strands, 2 and 5 (Figure 15.3B). Interestingly, the two classes of catalytic domain appear to have evolved in parallel and don’t segregate with the catalytic efficiency and accuracy, or type of DNA synthesis (replicative versus repair). Furthermore, reverse transcriptases (RT) and RNA-dependent RNA polymerases encoded by viruses possess pol I-like catalytic domains,29,34 and template-independent poly-A polymerases and CCA-adding enzymes are homologous to the pol b-like polymerases.35–37 RNA polymerases I, II and III, which transcribe DNA to RNA, share a conserved catalytic domain quite different from the DNA polymerases, CCA-adding enzymes, and poly-A and RNA-dependent RNA polymerases. The signature amino

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acid sequence of RNA polymerases, NADFDGD, is conserved among RNA polymerases from bacteria to high eukaryotes.38 This motif is located on a loop connecting two strands in a b-barrel domain, as observed in Taq and yeast RNA pol II38,39 (Figure 15.3C). This forms the third class of polymerase catalytic centres. Regardless of the differences in the tertiary structure and catalytic residue locations, all three classes of DNA and RNA polymerases appear to require two metal ions in the active site for substrate binding and the phosphoryl transfer reaction. In the best-studied DNA polymerase, pol b (X family), the two Mg2+ ions are jointly coordinated by two conserved Asps and the a-phosphate of an incoming dNTP30 (Figure 15.3B). Both Mg2+ are coordinated by six ligands in the octahedral geometry. One metal ion (conventionally known as the A metal ion) is close to the 3′-OH of the primer strand and suspected to activate it for nucleophilic attack, and the other metal ion (known as the B metal ion) is coordinated by the b and g phosphates (the leaving group) and is suspected to facilitate product formation. The metal ions are separated by ∼4 Å in the enzyme–substrate complex and by ∼3.4 Å in the enzyme transition-state complex.40 perhaps drawn close together by the transitional highly negatively charged phosphate anion. Perhaps not surprisingly, the same metal ion requirement and coordination geometry have been observed in pol l and m of the same X family30 and in the related CCA-adding enzymes.37 Interestingly, similar coordination of the two metal ions have been observed in pol I-like polymerases, including T7, Taq and Bst DNA pol of the A family,41–43 RB69 DNA pol of the B family,44 HIV RT,45 and Dpo4, pol k and Rev1 of the Y family.31 For the RNA pol II-like catalytic centre, in the currently available structures of RNA pol II complexed with a nucleic acid substrate, the metal ions are not coordinated as in DNA pol I or pol b, and are further apart than 4 Å. This may be due to the double conformations of the incoming UTP46 or the absence of the 3′-OH in the RNA strand.47 Given the three conserved Asp residues in the active site, it is likely that these carboxylates and the a-phosphate of the incoming NTP jointly coordinate the two Mg2+ ions for catalysis as with the other two classes of polymerases.

15.2.1 Alignment of Two Metal Ions and Fidelity of DNA Synthesis Displacement of two metal ions from the canonical binding sites in fact has been observed in the active site of several DNA polymerases. For example, Dpo4, which is capable of translesion synthesis, has a large, preformed and solvent-exposed active site.48 An omission of the 3′-OH in the primer strand, which is commonly used to crystallize polymerase–substrate ternary complexes, leads to ∼1 Å displacement of metal ion A.41,47,49,50 Although a variety of modified template bases (lesions) and mismatched replicating base pairs can bind in the active site of Dpo4,48 the catalytic efficiency differs with different substrates, and efficient nucleotide incorporation appears to correlate with the canonical configuration of two metal ions.50 Similar to Dpo4, DNA pol l also has a preformed active site, and one of the two metal ions is frequently missing from the active site in correlation with the misalignment of reaction groups in the crystal structures.51,52 Such correlation between the metal ion

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coordination and the preference of substrates may explain why polymerases exhibit strong substrate specificity that surpasses selection by base pairing. The rate-limiting step in polymerization is postulated to be the key step for rejecting a wrong incoming nucleotide and ensuring replication fidelity.53 A conformational step prior to the chemical reaction was previously suggested to be the rate-limiting step in DNA synthesis.54 Because of the structural observation of a large finger-domain movement induced by binding of a correct incoming nucleotide in Taq and T7 DNA pol and in HIV RT that transforms the active site from an ‘open’ to the ‘closed’ state,41,45,55 the fidelity of DNA polymerases was originally attributed to such an induced-fit conformational change. In recent years, this hypothesis has been challenged by presteady-state kinetic analyses of A, B and X family DNA polymerases.53 It has been shown that the finger domain movement or closing of the active site is much faster than the phosphoryl transfer reaction and therefore is not the rate-limiting step.56–60 Furthermore, analogous domain movement is not observed in all polymerases, for example, the X family pol l and all Y family members, which still exhibit respectable fidelity.27 These observations lead to the verdict that a step after dNTP binding and the finger-domain movement is the rate-limiting step. A rate-limiting conformational change that differentiates the correct versus incorrect incoming nucleotide has been observed for all polymerases, including Dpo4,61 whose active site doesn’t undergo finger-domain movement. The conformational step therefore has to be other than the domain movement and is likely subtle. The displacement of metal ions observed in the Dpo4–substrate complexes when the 3′-OH nucleophile is absent or the replicating base pair is a mismatched, suggests that binding of two Mg2+ ions in the active site is insufficient for nucleotide incorporation. The proper positioning and alignment of the two metal ions with regard to the catalytic residues and substrate may be the rate-limiting conformational changes and the key to converting small differences in binding energy to large differences in catalytic efficiency in all polymerases.

15.3 Nucleases That Require Two Mg2+ Ions in the Active Site RNase H and MutH represent two large superfamilies of nucleic acid enzymes that break backbone phosphodiester bonds using two-Mg2+ ion catalysis. The RNase Hlike superfamily includes the transposases and retroviral integrases with a DDE motif, argonaute endonuclease and the Holliday junction resolvase RuvC.12,62 The catalytic domains feature a central mixed b-sheet surrounded by a-helices (Figure 15.4A). Like DNA and RNA polymerases, two absolutely conserved Asps are the hallmark of the RNase H-like nucleases and transposases.63,64 They are located on two adjacent parallel b-strands, one long and one short (Figure 15.4A). One or two additional carboxylate residues (Asp or Glu) located on the surrounding secondary structures may be required for the catalysis. The MutH-like superfamily includes the type IIP restriction endonucleases (REases, see the comprehensive review65) and phage lambda Exo I.66 The catalytic centre marked by the DEK motif is located

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RNase H

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MutH

Figure 15.4 (Plate 16) Two families of nucleases that have been shown to use two-metal-ion catalysis: (A) RNase H is a prototype of the superfamily, which includes nucleases, retroviral integrases, RuvC Holliday junction resolvase and dicer argonaute. The two catalytically essential Asps are located on two adjacent parallel strands (highlighted in cyan). Two additional carboxylates may be required for catalysis. Two Mg2+ are jointly coordinated by the carboxylates and the scissile phosphate; (B) MutH represents another superfamily, which includes type IIP restriction endonucleases, T7 endonuclease I (a Holliday junction resolvasae) and T4 exonuclease I. The catalytic residues, Asp (D), Glu (E) and Lys (K) are located on the diverging point of a b-hairpin (highlighted in cyan). The loop that contains the first Asp, which plays the central role in Mg2+ coordination, is often disordered in the MutH-family nucleases in the absence of a properly aligned substrate or divalent cations. A diagram of the metal ion coordination in each case is shown below the actual structure (See colour plate section)

on the diverging point of a b-hairpin (Figure 15.4B). The single conserved Asp residue is located on the first strand of the hairpin. This strand is flexible and often disordered in the crystal structures. The remaining Glu–X–Lys (X is a hydrophobic residue) located on the second strand (Figure 15.4B) are not absolutely conserved. The Glu is occasionally substituted by Gln, and Lys by Glu or Gln.67 It has long been debated whether one, two or three metal ions are required for the phosphoryl transfer reaction catalysed by RNase H- and MutH-like nucleases and transposases. Disagreements typically stem from two general observations. Firstly, among the high-resolution structures of apo-RNase H, transposase and retroviral integrase, one or two metal binding sites have been identified and the locations of metal ions vary from structure to structure.68–71 Secondly, even in the presence

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of a DNA substrate, the number and location of metal ions vary among MutH-like type IIP REases,65 e.g. no metal ion in the Eco RI–substrate complex72 and three possible binding sites in Eco RV.73 Accompanying the metal ion variations, the active site composition of type IIP REases differs in the number of carboxylates required (two, three or four), and Lys of the DEK motif, catalytically essential for most of MutH-like endonucleases, is replaced by Glu in BamHI and Gln in BglII.67 These observations have raised speculation that REases may have different catalytic mechanisms using varying numbers of divalent cations.65,74 The crystal structures of a bacterial RNase H complexed with RNA/DNA hybrid substrates show two Mg2+ ions bound in the active site at a magnesium concentration below 2.5 mM.63 In the absence of a substrate, two Mn2+ ions were found at similar positions, but only at metal-ion concentrations much higher than are physiologically relevant.68 In the enzyme–substrate complexes, the two metal ions are jointly coordinated by the conserved Asp and scissile phosphate, each of which functions as a bidentate chelate (Figure 15.4A). By convention, the metal ion liganded to the nucleophile is called A, and the one liganded to the leaving group is B. A similar substrate-dependent two-metal-ion coordination is observed in crystal structures of the RNase H-like Tn5 transposase-DNA complexes.64 Among MutH-like nucleases, in the presence of their respective cognate substrates, MutH, BamHI and BglI bind two metal ions in the active site, in spite of the catalytic residue differences with Lys in MutH and BglI and Glu in BamHI.67 The coordination of the two metal ions by the scissile phosphate and a conserved Asp resembles the metal coordination observed in polymerases and RNase H (Figures 15.3 and 15.4), which is hereby referred to as canonical. It appears that in all cases at least one Asp is needed for coordination of the two metal ions. Asp is probably preferred because it has fewer rotamer conformations than Glu and costs less entropy to be fixed in one conformation. The metal ions observed in type IIP REases, PvuII, EcoRI and EcoRV, however, differ from the canonical ones in number and location.73,75,76 Such differences were originally attributed to different compositions of catalytic residues and perhaps different catalytic mechanisms. The conserved active site residues and tertiary structures of MutH, EcoRI PvuII and EcoRV suggest that these enzymes most probably share the same catalytic mechanism. The question is how to reconcile the metal ion differences observed in these related enzyme–substrate complexes.

15.3.1 Binding of Two Metal Ions in the Active Site Is Substrate Dependent Recently, crystal structures of human RNase H complexes with a RNA/DNA hybrid have provided new evidence on the substrate dependence of metal ion binding.77 One of the crystal forms contains 12 protein molecules in one asymmetric unit. Depending on the crystal lattice contacts, in some protein molecules the scissile phosphate is shifted away from the active residues by 1–2 Å, and these molecules contain only a single instead of two Ca2+ (Mg2+ mimics), and the Ca2+ binding site is neither the canonical A or B. This shows that not only are the number and location of divalent cations in the active site dependent on the presence of substrate,

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but they are also correlated to positioning of the scissile phosphate. Only when the scissile phosphate is appropriately bound in the active site are two divalent cations jointly coordinated by the phosphate and carboxylates for catalysis, as shown in Figure 15.4A. For type IIP REases, both the conformation of the active site residues and the binding of divalent cations depend on the DNA substrate. A cognate DNA sequence is required for two-metal-ion binding in the canonical positions, and, in return, divalent cations are required for high-affinity binding of the DNA substrate.78–80 Without divalent cations, EcoRV binds DNA with low sequence specificity, and in the presence Ca2+, which mimics Mg2+ for substrate binding, but inhibits hydrolysis, EcoRV binds the cognate recognition sequence specifically.81 Structural investigation revealed that when BamHI is associated with the sequence GAATCC (one base pair substitution of the cognate GGATCC sequence), although the binding is tight enough for crystallization, the DNA is shifted away from the binding site of the cognate sequence and metal ions are absent.78 Moreover, the conserved Asp,65 which coordinates both metal ions in MutH, BamHI and BglI, is located on a loop that is disordered in the absence of a cognate DNA, partially ordered in the presence of DNA, and fully ordered only when both metal ions are bound.67,78 Without divalent cations, the catalytic residues of EcoRI in all DNA-complex structures, although ordered, are oriented in such a way that they are incapable of catalysis.72 MutH cleaves the unmethylated strand in the hemimethylated GATC sequence 10–20-fold better than the unmethylated duplex. In the crystal structures of the MutH–DNA complexes, the active site is clearly better ordered and better oriented for catalysis with the hemimethylated GATC than the unmethylated.67 Both the active site conformation and the metal ion binding in MutH-like nucleases are consistently shown to depend on binding of a cognate DNA substrate. The dependence of the nucleases on the cognate DNA substrate for metal-ion binding provides an explanation as to why the number and location of metal ions in the structures of homologous type IIP REases vary. To crystallize nuclease– substrate complexes, cleavage has to be prevented by artificial means, for example, substitution of catalytic residues, use of inert substrate analogues, or divalent cations that do not support catalysis. Strategies used to capture enzyme–substrate complexes with the purpose of preventing chemistry from taking place inevitably perturb the protein–DNA interactions. In addition, crystal lattice contacts may distort macromolecular interactions. One has to interpret crystal structures and particularly the active site conformation, including the number and location of metal ions, with caution. The exquisite sensitivity of metal ions to alterations in coordination environment probably underlies the specificity of these enzymes.

15.3.2 Two Metal Ions Enhance Catalytic Specificity When two divalent cations were first observed in the active site of alkaline phosphatase and the exonuclease of the Klenow fragment,82,83 they appeared to mediate binding of nucleic acids in the carboxylate-rich active site. The two metal ions are located roughly in parallel to the phosphosugar backbone on the opposite side of

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Figure 15.5 Alignment of metal ions in the two-metal-ion mechanism. Two metal ions, preferably Mg2+, are jointly coordinated by at least one absolutely conserved Asp and the scissile phosphate, which is the a-phosphate of (d)NTP in the case of polymerase. Metal ion A is directly coordinated to the nucleophile and metal ion B to the leaving group. The reaction products are 3′-OH and 5′-PO4. (Reproduced from Yang, W. et al., Making and breaking nucleic acids: two-Mg2+-ion catalysis and substrate specificity, Mol. Cell, 2006, 22, 5–13)

the bases and bisected by the plane of the trigonal core of the pentacovalent intermediate (Figure 15.5). Based on the particular coordination geometry of the two metal ions, Steitz and Steitz proposed a generally applicable two-metal-ion mechanism, by which metal ion A acts like a Lewis base to reduce the pKa of a nucleophile and facilitate its deprotonation and metal ion B stabilizes the pentacovalent intermediate and the 3′ oxyanion leaving group.84 In the last 14 years, many structures of polymerases and nucleases using two-metal-ion catalysis have been determined.1,29–31 In addition to facilitating catalysis, the metal ions appear to play a critical role in enhancing substrate recognition and catalytic specificity for both polymerases and type IIP REases. By being sensitive to the coordination environment, the metal ions induce misalignment between a nonpreferential substrate and catalytic residues, thus greatly reducing the catalytic efficiency. Catalysis occurs efficiently only with a preferred substrate. The enhancement of substrate recognition is not limited to the fidelity of DNA and RNA polymerases, and sequence and methylation specific nucleases, but also extends to structure-specific nucleases, e.g. RNase H and argonaute. Although the latter nucleases have no explicit sequence specificity, RNase H only cleaves the RNA strand in a RNA/DNA hybrid and not in dsRNA, while argonaute functions in tandem with a duplex ruler and generates cleavage products of a specific length.85

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15.3.3 Movement of the Two Metal Ions During Transition State Formation In all polymerases and nucleases reported to date, the two metal ions are jointly coordinated by a conserved Asp and the scissile phosphate (Figures 15.3 and 15.4), and their general positions conform to the 1993 proposal84 (Figure 15.5). The precise position and separation distance between the two metal ions, however, may vary at each reaction step according to the changes in the coordination environment. Tuning of the two metal ions is evident in both DNA pol b and RNase H. In the crystal structure of pol b–substrate complexes, the two metal ions are 4.0 Å apart, and in the structure of pol b-transition state-like complexes, they become 3.4 Å apart.40 Similarly, the separation of the two Mg2+ ions in the RNase H–substrate complexes vary from 4.0 Å to 4.5 Å, depending on which one of the catalytically essential carboxylates was replaced by carboxylamide.63 When the separation is 4.5 Å, the mutant protein (D132N) is completely inactive. But the mutation (D192), in which the metal ions are 4.0 Å apart, has detectable nuclease activity in the presence of Mn2+.63 The 4.0 Å separation allows the two metal ions to fit snugly on each side of the scissile phosphate, as depicted by Steitz and Steitz84 (Figure 15.5). However, the potential nucleophile coordinated by metal ion A is 3.5 Å or further away from the target phosphorus atom in the nuclease–substrate complexes (Figure 15.5). For nucleophilic attack to occur, the hydroxide ion probably needs to be within 2.5 Å of the phosphorus. The nucleophile is unlikely to dissociate from metal ion A and move over 1 Å by itself, owing to the high energy cost of removing an inner-shell metal-ion ligand and the repulsion between the negatively charged nucleophile and the scissile phosphate. We hypothesize that the 4.0 Å separation of the two metal ions represents a ‘resting’ state, and that metal ion A moves towards metal ion B and brings the nucleophile within striking distance for phosphoryl bond formation. Bringing the two metal ions closer could also better neutralize the developing negative charge on the pentacovalent intermediate. A 3.5 Å separation between two Mg2+ ions has been observed with DNA pol b and RNase H, when complexed with transition state mimics.86 Interestingly, such close juxtaposing is not observed when an Asp coordinating both metal ions is replaced by Asn (D132N).86 In addition, two Mg2+ ions are reported to be as close as 3 Å in the T7 RNA polymerase–substrate complex structure determined at 2.88 Å49 (Figure 15.3A). It is possible to bring two Mg2+ ions closer than 4.0 Å because its interatomic distance is 3.197 Å.87 Ca2+ has been widely used to substitute for Mg2+ during enzyme–substrate complex formation because it usually supports substrate binding. Catalysis of all polymerases and the nucleases mentioned in this chapter is inhibited by Ca2+.67,88–90 Comparisons of crystal structures of nucleases and polymerases reveal little difference between Ca2+ and Mg2+ coordination geometries.67,91,92 Interestingly, the interatomic distance of Ca is 3.94 Å,87 and the closest approach between two Ca2+ ions observed to date is 3.8 Å.93,94 This could be one reason that Ca2+ is unable to support catalysis of two-metal-ion-dependent phosphoryl transfer reactions. In contrast, the size and charge density of Zn2+, Co2+, Cd2+ and Mn2+ are comparable to Mg2+, and these divalent cations can substitute for Mg2+ in catalysis.

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15.3.4 Metal Ions A and B Together Destabilize the Substrate and Assist Product Formation Metal ion A has been suggested to play an essential role in nucleophile formation in ribozymes and enzymes that lack a conventional general base.84 Experimental observations accumulated over the years suggest that nucleophile formation cannot be achieved by metal ion A alone. In the absence of an apparent general base, a potential nucleophile is often coordinated by the nucleic acid substrate, in addition to metal ion A. For example, in RNase H, the potential nucleophilic water molecule interacts with metal ion A and the pro-Rp nonbridging oxygen of the phosphate 3′ to the scissile bond,63 and in MutH, a Lys sidechain supplies the third ligand67 (Figure 15.4B). Neither phosphate oxygen nor Lys is a conventional general base. Interestingly, although the pro-Rp oxygen coordinates the nucleophile in both RNase H and MutH, its stereospecific replacement with sulfur has the opposite effect. The Rp sulfur enhances the cleavage activity of MutH,67 but it inhibits RNase H.95 MutH, whose active site contains a Lys, likely benefits from the extra negative charge brought by the Rp sulfur (due to the nature of the single P–S).96 But RNase H, whose active site contains four carboxylates, may be inhibited by the extra negative charge and steric hindrance due to the sulfur substitution (larger atomic radius than oxygen). Parallel to the diverse ways of substrate-assisted nucleophile formation, the coordinating carboxylate for metal ion A can be replaced by carboxylamide or histidine in the RNase H family.63,97 It appears that the net charge environment contributed by both the catalytic residues and the substrate is the key for twometal-ion catalysis. Metal ion B may promote the phosphoryl transfer reaction instead of passively stabilizing the transition state, as originally proposed.84 Even though the A metal ion seemingly initiates the chemical reaction by directly coordinating to the nucleophile, replacement of carboxylates that coordinate metal ion A often do not eliminate nuclease activities as effectively as mutating the B metal ion ligands.63,98 In addition, the B site also exhibits a stronger metal ion preference than the A site. In the 3′ to 5′ exonuclease of DNA polymerase I, the B site normally binds a Mg2+ ion,99 but it rejects Mg2+ when the nonbridging oxygen of the scissile phosphate is replaced by sulfur.100 In the crystal structures of RNase H–substrate complexes, the Mg2+ ion at the B site is coordinated irregularly by five ligands, none of which is a water molecule (Figure 15.6A).63 The nonideal coordination geometry and ligands devoid of water suggest that metal ion B in the enzyme–substrate complex is in a high-energy state and may strain the scissile phosphodiester bond. The deviation from ideal octahedral coordination geometry may explain why Mg2+ is preferred over Ca2+ for catalysis. Ca2+ readily accepts six, seven, eight or nine ligands in various coordination geometries, and promiscuity regarding ligand type and coordination geometry renders Ca2+ less likely to destabilize the enzyme–substrate complex and promote product formation. This may be another reason that Ca2+ does not usually replace Mg2+ in phosphoryl transfer reactions. In a mutant RNase H–product complex, the B-site Mg2+ shifts by 0.9 Å and becomes coordinated in near octahedral geometry by six ligands, two of which are water molecules (Figure 15.6B).86 The change from five-ligand coordination of metal

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Figure 15.6 (Plate 17) Coordination of the two metal ions changes during RNA cleavage by RNase H: (A) Enzyme–substrate complex; (B) Enzyme–product complex (See colour plate section)

ion B in the substrate complex to a favourable six-ligand coordination in the product complex indicates that the two metal ions also play a role in product formation. The two metal ions jointly coordinated by the scissile phosphate may also facilitate product release. After splitting into 5′-phosphate and 3′-OH, the cleaved scissile phosphate can no longer coordinate the two metal ions simultaneously. At least one metal ion has to be displaced or released, as observed in product complexes of DNA and RNA polymerases and nucleases.49,50,73,86,91 The RNase H–product complexes have been trapped in the crystal form by a key residue for product release (E188) being substituted by Ala. In the product complex the cleaved scissile phosphate moves out of the active site, and concomitantly metal ion A moves by 2 Å and retains only one ligand from RNase H.86 Presumably eventual dissociation of the metal ion leads to the release of the cleavage product. Instead of having separate responsibilities, as initially proposed, the two metal ions appear to work together throughout phosphoryl transfer reactions. It is widely accepted that enzymes facilitate chemical reactions by proper alignment of substrates relative to each other and to the catalytic residues, thereby lowering the energy barrier between the substrate and product states. As such, the key element of two-metal-ion catalysis may be the proper alignment of the two metal ions with regard to the conserved carboxylates and substrate. The strong conservation of twometal-ion coordination in polymerases, nucleases and group I intron (in which a backbone phosphate replaces the conserved Asp to coordinate both metal ions)101 certainly supports this metal-ion centric view. Once the proper alignment is formed and the enzyme–substrate complex is destabilized, a nucleophile can form and be activated in various ways without a conventional general base. Similarly, after the reaction proceeds past the pentacovalent intermediate, protonation of a leaving group is energetically favourable, even without a general acid. This view is supported by the biochemical and computational studies of metal-ion-dependent proton transfer in DNA and RNA polymerases.102–104

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15.4 Advantages of Two-Metal-Ion Catalysis: Specificity and Versatility The exquisite sensitivity of metal ions to the local environment16,18,105 provides a means for nucleic acid enzymes to convert small conformational differences into large catalytic rate changes. Three features of two-metal-ion mechanisms ensure the substrate recognition and catalytic specificity: (i) interdependence of substrate recognition and two-metal-ion binding, as illustrated in RNase H,63 type II restriction endonucleases,67,78,81 ribozyme106 and DNA polymerases50,107; (ii) usage of metal ions with highly stringent coordination requirements to destabilize the enzyme–substrate complex and promote product formation, as observed with RNase H86 and (iii) reliance on the electrostatic environment created by both the catalytic residues and the nucleic acid substrate for nucleophile formation, and for metal ion movement during the course of the phosphoryl transfer reaction. Mg2+, known for its exquisite octahedral coordination geometry and narrow distribution of coordination distance,18 is most often used in two-metal catalysis. Ca2+, because of its large atomic radius and indifference to ligand number and coordination distance, usually does not support phosphoryl transfer reactions. Mn2+, which is most similar to Mg2+ in chemical nature, but has relaxed coordination requirements, can replace Mg2+ and allow phosphoryl transfer reactions to occur with altered catalytic residues, nonideal substrates and reduced specificity.50,108–112 The natural abundance of cellular Mg2+ over Mn2+ thus furthers the substrate specificity of two-metal-ion catalysis. The second advantage of two-metal-ion catalysis is its versatility and reversibility. The versatility is clearly demonstrated in DNA and RNA synthesis, where polymerases have to incorporate various nucleotide building blocks ((d)NTPs) with roughly equal efficiency and yet in each reaction cycle select only the right one according to the template base. Two-metal-ion catalysis, which allows the active site to align correctly only upon association with a Watson–Crick base pair, enables DNA and RNA polymerases to achieve high specificity with a broad range of substrates. Regarding reversibility, since two-metal-ion catalysis has no specific requirement for a general base or acid, it supports both making and breaking of DNA and RNA. During DNA transposition and RNA splicing, a single active site with two metal ions can carry out at least two consecutive phosphoryl transfer reactions.63,84 In the first reaction, DNA or RNA is cleaved to generate a 3′-OH (product), and in the second reaction the 3′-OH is the nucleophile (substrate) and results in strand transfer (Figure 15.1). The symmetric arrangement of the two metal ions in the active site of Tn5 transposase and group I ribozyme (Figure 15.7) allows the metal ions to alternate their roles in activating a nucleophile and stabilizing a pentacovalent intermediate in consecutive phosphoryl transfer reactions without releasing the 3′-OH.64,101 15.4.1 One-Metal-Ion Catalysis Exceptions to two-metal-ion catalysis are one-metal-ion and metal-ion-independent catalysis utilized by certain nucleases and recombinases. To date, only reactions resulting in 5′-OH and 3′-phosphate are known to be catalysed by enzymes that

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Figure 15.7 (Plate 18) Coordination of the two metal ions in Group I intron. The two metal ions have similar coordination geometry (hence symmetric) and can reverse their roles as ‘A’ and ‘B’ metal ions in the successive steps of RNA splicing reaction. (Reproduced from Yang, W. et al., Making and breaking nucleic acids: two-Mg2+-ion catalysis and substrate specificity, Mol. Cell, 2006, 22, 5–13) (See colour plate section)

don’t require divalent cations, for example, hepatitis virus delta ribozyme,113 RNase A, type IB topoisomerase, phage l integrase and g d resolvase.114 Reactions resulting in 5′-phosphate and 3′-OH usually require one or two metal ions. Nucleases with the bba-metal motif, for example His-Cys and CIY-YIC homing and HNH endonucleases74,115–118 have no conserved catalytically essential carboxylates in the active site and require one single divalent cation for catalysis. The single metal ion occupies the same position relative to the scissile phosphate as metal ion B of the twometal-ion mechanism. These nucleases often depend on a conserved His to activate a nucleophile. Without the requirement for specifically aligning two metal ions, a variety of divalent cations are able to support one-metal-ion catalysis.118 These types of nucleases are either sequence nonspecific or specific. If sequence specific, the cleavage and binding specificity are equal and specificity is achieved by noncatalytic domains that recognize 20–40 bps, much longer than the 4–8 bps typically recognized by type II REases.119 15.4.2 Prediction of Two-Metal-Ion Catalysis Two-metal-ion catalysis, because of its unique specificity and versatility, can be readily predicted based on the composition of catalytic residues, the nature of

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nucleophile and substrate specificity. If all, or the majority of, the catalytically essential residues are carboxylates, with at least one invariable Asp, and if the enzyme exhibits stricter catalytic specificity than binding specificity, catalysis almost certainly depends on two metal ions. Structure-specific FEN-1 and XPF endonucleases,120,121 RNase III and dicer endonucleases,97,122 and Group II ribozymes catalysing RNA splicing with high substrate specificity, clearly fulfil the criteria for a twometal-ion catalysis. A noted variation of two-metal-ion catalysis is the three metal ions found in homodimeric LAGLIDADG homing endonucleases,123 where a single metal ion B is shared between two neighbouring active sites. Nevertheless, each phosphoryl transfer reaction requires two metal ions coordinated by a conserved Asp.

15.5 Concluding Remarks In this chapter we have summarized the following properties of two-metal-ion catalysis: (i) participation of the two metal ions in substrate recognition; (ii) the role of metal ions, particularly at the B site, in destabilizing the enzyme–substrate complex and driving the reaction forward; (iii) dependence on a specific electrostatic environment rather than metal ion A per se for the nucleophile formation and nucleophilic attack; (iv) movement of two metal ions towards each other to stabilize transition state and facilitate product formation; (v) movement of two metal ions away from each other to release products. The key to two-metal-ion catalysis is the proper alignment of the metal ions with regard to the catalytic residues and the nucleic acid substrate. The extensive involvement of the nucleic acid substrate in metal-ion binding and the sensitivity of metal ions to the coordination environment enable nucleic acid enzymes that utilize two-metal-ion catalysis to convert small thermodynamic differences into large changes in catalytic rate and achieve extraordinary specificity.

Acknowledgements I thank Dr R. Craigie for reviewing the manuscript and the Intramural Research Program of the NIH, NIDDK for funding the research. Figures 15.1A, 15.5 and 15.7 are reproductions from Mol. Cell, 22, 5–13 Figures 2A, 1B and 3C, respectively.

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Part D Toxicological Aspects

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

16 Structural Studies on the MercuryII-Mediated T-T Base-Pair Using NMR Spectroscopy Yoshiyuki Tanaka and Akira Ono

16.1 The History of the MercuryII-Mediated Thymine-Thymine Base Pair Studies on the mercuryII–DNA interaction began with the paper by Katz in 1952.1 In that report, Katz performed light scattering and viscosity measurements on calf thymus DNA in a complex with mercury ions (HgII), and found that HgII can bind to DNA in a reversible manner. In 1954, Thomas followed Katz’s experiments with HgII titration experiments on calf thymus DNA using UV spectra.2 Yamane, Davidson and Dove performed extensive HgII titration experiments with UV spectra, using E. coli DNA3,4(1960, 1961), M. lysodeikticus DNA4(1961), poly-d(AT)4(1961), poly-rA5(1962), poly-U5(1962) and tobacco mosaic virus RNA5(1962). The results they obtained from these studies showed: (i) upon one HgII binding to DNA, two protons are released in the initial stage of HgII-titrations; (ii) in the HgII titration experiments of natural DNA (E. coli and M. lysodeikticus DNA), UV spectra showed an isosbestic point until the molar ratio of [HgII]/[mole of nucleotides of the DNA] became 0.5; (iii) in the case of poly-d(AT) (adenine (A)–thymine (T) alternating polymer), no isosbestic point was observed for the UV spectra of HgII titrations; (iv) in the case of poly-d(AT), the number of released protons upon HgII-binding (∆[H+]/∆[HgII]) decreased at a molar ratio ([HgII]/[mole of nucleotides of the DNA]) of 0.25; (v) in the HgII titration of poly-uridine(U), UV spectra showed an isosbestic point until the molar ratio of [HgII]/[mole of Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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R

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O

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Scheme 16.1 O H II N N Hg

N R

N N

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O

N R

Scheme 16.2 nucleotides of the DNA] became 0.5; (vi) a possible HgII binding site might be N3 of T/U. Based on the above results, Katz reconsidered the stoichiometry of HgII binding to DNA molecules6,7(1962), and proposed a chemical structure of the HgII mediated T-T base pair (T-HgII-T) as a possible interaction mode in DNA molecules8(1963) (Scheme 16.1). Furthermore, he presented a possible mechanism for DNA strand displacements based on the T-HgII-T pair. To obtain quantitative data, thermodynamic studies on HgII–nucleoside complexes were performed by Eichhorn and Clark9(1963), and Simpson10(1964). Simpson determined association constants of HgII to nucleosides under various pH values. In 1965, Nandi et al. proposed a unique interaction mode for an A-HgII-T base pair that is different from the T-HgII-T base pair11 (Scheme 16.2). In 1966, Gruenwedel and Davidson reported UV-monitored titration experiments on natural DNA molecules (calf thymus DNA, bacteriophage T4 DNA and M. lysodeikticus DNA) with methymercuric hydroxide (CH3HgOH).12 Based on the Katz model of the T-HgII-T pair, they porposed that methymercuric ions can denature DNA molecules by HgII binding to N3 of T and N1 of guanosine (G). These pioneering works stimulated many structural studies of the HgII-mediated T-T base pair in DNA duplexes, with crystallography, vibrational spectroscopy and NMR spectroscopy being applied.

16.2 Crystallographic Studies on HgII–Nucleobase Complexes There are two kinds of crystal structures for an HgII–complex with uracil (U) or thymine (T) (Figure 16.1). In 1971, Carrabine and Sundaralingam determined the structure of a HgII–uracil (U) complex13 (Figure 16.1a). In this structure, two uracil molecules are bound to HgII at their O4-carbonyl oxygen [O4(U)]; this structure was different, however, from the putative structure of the T-HgII-T pair proposed by Katz. In 1974, Kosturko, Folzer and Stewart reported a crystal structure of the

UV, UVCD and Vibrational Spectral Studies 441 (a)

H O

O

N N H

O

HgII

H N

O N

H

(b)

O N

N O

O HgII

N

N

O

Figure 16.1 Experimentally observed U-HgII-U13 and T-HgII-T14 base pairs in crystals. Solution conditions for crystallization: (a) 9.72 mM uracil and 4.86 mM HgCl2 at pH 4.2; (b) 20 mM 1-methylthymine and 10 mM HgO

1-methylthymine–HgII (2 : 1) complex14 (Figure 16.1b). Interestingly, this structure essentially reflects Katz’s model. This crystal structure demonstrated for the first time that N3 of T can also be an HgII binding site. Although these two crystal structures do not directly indicate that the binding modes observed in the crystals correspond to those observed in natural DNA, the determined structures were important experimental data for considering an HgII binding mode. On the other hand, there are studies exploring other types of HgII–U complexes and HgII complexes with other nucleobases15–28 (Figure 16.2). Beauchamp and coworkers reported crystal structures of methylmercury complexes with adenine and cytidine (Figures 16.2a–g).15–19 In 1988, Sheldrick and Gross also reported methylmercury complexes with 7-methylguanine (Figures 16.2h,i).20 Lippert and coworkers reported HgII complexes with 1,3-dimethyluracil at the C5-position (Figures 16.2j– l),23–25 an HgII complex with 9-methyladenine at the N7-position (Figure 16.2m)26 and bimetallic HgII,PtII complexes with 1-methylcytosine (Figures 16.2n,o).21,27 These data indicated the intrinsic potential of nucleobases to form various types of metal complexes.

16.3 UV, UVCD and Vibrational (IR/Raman) Spectral Studies As mentioned above, UV absorption spectra were employed from the early stages of studies on the HgII–DNA interaction. UV spectral studies12,29 were continued by Gruenwedel’s group, which developed their study into UVCD spectra.30–32 They presented comprehensive UVCD spectra of HgII–DNA complexes under various conditions (see references for details). In the case of vibrational (IR/Raman) spectra, complexations between methylmercuric ions and nucleosides were studied (Figure 16.3).33–44 In 1974, Mansy, Tobias and coworkers reported Raman spectra of a CH3HgII–uracil complex with an N3-HgII bond (Figure 16.3a),33 a CH3HgII–5′-GMP (guanosine-5′monophosphate) complex with an N1-HgII bond at neutral pH in H2O solution (Figure 16.3b)34 and a CH3HgII–5′-GMP complex with an N7-HgII bond at acidic pH in H2O solution (Figure 16.3c).34 Proton–HgII exchange processes are required to form the complexes (Figures 16.3a,b), and the derived Raman spectra provide reference data for the T-HgII-T base pair. There are also Raman spectra of CH3HgII–cytidine and CH3HgII–5′-AMP (adenosine-5′-monophosphate) complexes in H2O solution (pH 3, 5 and 7) (Figures 16.3d,e),36 the HgII–uracil complex with O4-HgII

442

Structural Studies on the MercuryII-Mediated T-T Base Pair

Figure 16.2 Experimentally observed HgII complexes in crystals. References for the crystal structures and crystallization conditions are as follows: (a) and (b): 15; (c): 16,17; (d): 18; (e), (f) and (g): 19; (h) and (i): 20; (j): 24; (k): 23; (l): 25; (m): 26; (n): 21; (o): 27; (p): 22. Solution conditions for crystallization: (a) aqueous mixture of CH3HgNO3 (1 M aqueous solution) and adenine (67–100 mM aqueous solution) in 1 : 1 ratio; (b) aqueous mixture of CH3HgClO4 (1 M aqueous solution), NaOH (1 M aqueous solution) and 9-methyladenine in 2 : 1 : 1 ratio; (c) crystallization details are not given; (d) aqueous mixture of CH3HgNO3 or CH3HgClO4 (1 M aqueous solution) and adenine in 1 : 1 ratio; (e) CH3HgOH, CH3HgNO3 (∼0.5 M aqueous solution) and 1-methylcytosine (methanol) at 2 : 1 : 1 ratio; (f) CH3HgOH (1 M aqueous solution) and 9-methyladenine (acetonitrile) at 2 : 1 ratio; (g) aqueous mixture of CH3HgNO3 or CH3HgClO4 (1 M aqueous solution) and adenine (67–100 mM aqueous solution) in 3 : 1 ratio; (h) 54 mM CH3HgOH and 54 mM 7-methylguanine at pH 9–12; (i) 54 mM CH3HgOH and 18 mM 7-methylguanine at pH 1–3; (j) 18 mM Hg(O2CCH3)3 and 18 mM 1,3-dimethyluracil at pH 3.5; (k) 20 mM Hg(O2CCH3)(1,3-dimethyluracil-C5) and 20 mM KCN in water; (l) 47 mM Hg(O2CCH3)(1,3-dimethyluracil-C5) and 47 mM 9methyladenine at pH 1.2; (m) 44 mM Hg(NO3)2 and 88 mM 9-methyladenine at pH 2.5; (n) 6.8 mM HgO and 6.8 mM trans-[Pt(NH3)2(1-methylcytosine)2](NO2)2 at pH 5.8; (o) 9.9 mM Hg(O2CCF3)2 and 10 mM trans-[Pt(NH3)2(1-methylcytosine-N4)2](NO2)2, raising pH 1.8 to 2.9

bonds in the crystal phase (Figure 16.3f),37 the HgII–5′-UTP (uridine-5′triphosphate) complex with C5-HgII bonds in H2O solution (Figure 16.3g)37 and CH3HgII–inosine complexes with N1-HgII bond (pH 8) and an N7-HgII bond (pH < 2) in H2O solution (Figures 3h,i).38Also,Raman spectra of a CH3HgII–5′-TMP (thymidine-5′monophosphate) complex in H2O (Figure 16.3j), complexes between CH3HgII and 5′-TMP, 5′-AMP, 5′-GMP and 5′-CMP (cytidine-5′-monophosphate) mixtures in

UV, UVCD and Vibrational Spectral Studies 443

Figure 16.3 Compounds for which Raman/FTIR spectra were recorded. References for these compounds are as follows. (a): 33; (b) and (c): 34; (d) and (e): 36; (f) and (g): 37; (h) and (i): 38; (j), (k) and (l): 39; (m): 39,42; (n), (o), (p) and (q): 40; (r), (s) and (t): 41; (u): 39; (v) 43

444

Structural Studies on the MercuryII-Mediated T-T Base Pair

H2O (Figure 16.3k), a CH3HgII–1-methylthymine complex in H2O (Figure 16.3l) and a crystalline HgII–1-methylthymine complex (Figure 16.3m) were presented.39 In addition, they presented IR spectra from solid samples of organomercury–guanosine/ thymidine complexes (Figures 16.3n–q).40 In 1982, Beauchamp, Savoie and coworkers also reported IR/Raman spectra of methylmercury–adenine complexes (Figures 16.3r–t).41 Interestingly, in 1977, Mansy, Tobias and coworkers recorded Raman spectra of calf thymus DNA in the presence of CH3HgIIOH (Figure 16.3u).39 This data provided Raman spectroscopic information on HgII and natural DNA molecules. As mentioned in Section 16.2, there are three types of HgII–T/U complexes, namely N3-, O4- and C5-mercurated ones. In the above noted vibrational studies, all types of complexes in water solutions were characterized by Raman spectra and several marker bands for detecting each HgII–U/T complex were found. Notably, not only solution samples, but also crystal samples prepared by means of Kosturko’s method14 (with N3-HgII bonds) and Carrabine’s method13(with O4-HgII bonds) were employed. This fact implied that the derived FT-IR/Raman spectra surely originate from the crystal structure and are not from any other structure. Therefore, the data obtained from the studies provides rigid and reliable basal data for the assignment of Raman bands from other systems. In 2001, Morzyk-Ociepa and Michalska reported very fine FT-IR and Raman spectra of solid samples of HgII complexes with 1-methylthymine.42 Furthermore, they performed quantum chemical calculations of vibrational spectra, and extensively assigned IR/Raman bands.42 In 2003, they reported FTIR and FT-Raman spectra of AgI-1-methyluracil complexes (U-AgI-U complex) (Figure 16.3v) that could be used as reference data for the T-HgII-T base pair.43 In 2004, they recalculated the theoretical frequency of vibrational spectra, and refined the assignments of Raman and IR bands.44 Therefore, these data can be applied to the interpretation of FT-IR/Raman spectra in relation to specific vibrational modes. For vibrational spectroscopy, sufficient amounts of accurate spectral data of HgII–nucleoside complexes have been collected, along with data on neutral thymine.45 Especially, in terms of the structural studies on T-HgII-T base pairs, HgII–ligand interactions within THgII-T base pairs can be precisely analysed.

16.4 NMR Spectral Studies 16.4.1 Nucleoside–Metal Ion Systems NMR studies on nucleobase–metal ion(ZnII) interactions in dimethyl sulfoxide (DMSO) began in the 1960s.46,47 These studies extended to nucleoside–HgII interactions.48,49 At that time, NMR chemical shifts and their perturbations upon metal salt titrations were used to understand the nucleoside–metal cation/counteranion interactions.46–52 In 1975, Sohma and coworkers extensively studied chemical shift perturbations of 1H and 13C nuclei in cytidine upon the addition of various metal cations and counteranions to study how they affect 1H and 13C chemical shifts.52

NMR Spectral Studies

445

In 1978, Marzilli and coworkers monitored the HgII binding to cytidine using C NMR spectroscopy. In their report, higher-field shifts of carbon atoms (C2, C4) were reported (Table 16.1).53 Next, they reported 1H and 13C NMR spectra of complexes between HgII and nucleosides (uridine, cytosine and adenosine) in dimethylsulfoxide (DMSO).54,55 In that study, they found that cytidine bound to HgII, probably at N3, but that uridine did not bind to HgII through its N3 atom in DMSO.54,55 This observation was somewhat controversial, since proton–HgII exchange reactions in H2O had already been proposed for uridine. In 1982, however, they found that proton–HgII exchange reactions in DMSO can occur at the N1 of guanosine ‘in the presence of triethylamine base’ (Table 16.1).56 Through the 13C NMR experiments, moderate but significant chemical shift perturbations were observed for the resonances from the carbon atoms adjacent to the metal-binding nitrogen atoms (Table 16.1). For the HgII–thymidine system, Buncel and coworkers in 1981, 1985 and 1986 reported 13C NMR spectra of the complexes (Table 16.1).57–59 In these reports, they isolated the complexes of HgII–thymidine and HgII–guanosine, and dissolved these complexes in DMSO.57–61 The derived solutions were used for NMR measurements. According to these data, significant lower-field shifts of the 13C nuclei adjacent to HgII-binding sites (N3) were observed for C2 and C4 nuclei (Table 16.1) (see references for details).58,59 These data strongly supported the HgII-T complexation through N3, as proposed by Katz.8 Interestingly, Buncel and coworkers reported that proton– HgII exchange reaction occur at guanine to give a HgII–guanosine covalent complex through N1. This is because they observed the lower-field shifts of the 13C chemical shifts of C2 and C6 (N1-neighbours) (see references for details).57–59 In 1982, 1983 and 1986, Buchanan and coworkers monitored HgII binding processes to guanosine, cytosine, adenosine and inosine in DMSO by using 15N NMR spectroscopy (Table 16.2).62–64 From their experiments, very large chemical shift perturbations were observed for N7 of guanosine (∼20 ppm higher field), which meant that the HgII binding site in guanosine was not N1 but N7 (Table 16.2)62 Under their conditions, proton–HgII exchange did not occur. However, this appeared to be because NMR spectra were recorded in the DMSO solution without any base, which is indispensable for a deprotonative metallization of guanosine at N1 in DMSO, based on the observations by Marzilli and coworkers.56 Next, Buchanan and coworkers studied the methylmercuryII complexes with 5′-GMP and 3′-CMP using 15 N NMR spectroscopy.64 It was pointed out that at pH 8 in H2O, HgII binding sites became the N1 of 5′-GMP with proton–HgII exchange processes (Table 16.2).64 In another water solution system, Polak and Plavec, in 1999, reported 5N NMR spectra of HgII complexes with methyl esters of 5′-GMP and 3′-GMP (guanosine3′-monophosphate) (Table 16.2).65 In their report, both compounds were seen to bind to HgII through their N7 atoms. They postulated that chelation of HgII with N7 and the phosphate group makes HgII bind to N7. With these data, however, HgII binding sites of various nucleosides were determined with 15N NMR spectroscopy. In a later section (Section 16.4.2), 15N NMR data on the proton–HgII exchange systems for a DNA duplex are described. In respect of another interacting point, 199Hg NMR spectra of HgII–nucleoside complexes were recorded by Norris and Kumar, in 1984.66,67 They reported 199Hg 13

Hg (0.5 M) CH3HgII (0.2 eq.)

−2.2 ∼−0.5

N3 −2.7 – ∼0.7

−1.24 – −0.66 −1.25

+0.5 +0.3 +0.9 +1.1

−0.2 +0.5 +0.1 +0.9

−1.8

−0.7

+1.7 ∼+1.5

–

– – – – –

−2.63 −2.76 −1.27 −2.60

n.a. n.a.

n.a. n.a. n.a. n.a. – – – –

n.a. n.a. n.a. n.a.

+0.5 n.a. −1.78 n.a. – n.a. −0.37 n.a. −1.42 n.a. +2.1 n.a. +0.2 n.a. n.a. ∼+1

5-CH3

C5

+0.4 +1.04 – +0.24 +0.98 −0.5 −2.0 ∼0

n.a. n.a. n.a. n.a.

C8

+0.9 ∼−0.2

n.a. n.a.

+0.13 +0.07 +0.48 +0.32 +2.5 – +2.3 +1.1

−1.04 +1.61 +1.75 −0.37 +0.67 −0.85 +1.88 −0.03 N.D. −0.11 −0.04 – – – –

–

+4.6 +2.75 +3.04 +2.58 −0.40 +5.6 +3.4 ∼+3

0.0 −0.3 +0.4 +0.3

C6

53 64

65 65 65 65 103 103 76, (105) 104

56 56 56 56

58, (59) 56 56 56 56 58, (57) 57 64

58, (59) 58 80 80

Reference

a Chemical shift changes are listed in ppm. Positive and negative values represent lower-field shifts and higher-field shifts, respectively. MepdG: methylester of 5′-GMP; dGpMe: methylester of 3′-GMP; n.a.: not applicable; N.D.: not detected; –: not recorded. b For DNA/RNA oligomers, the chemical shift changes of the underlined residues are indicated.

cytidine (0.2 M)/DMSO/3′-CMP/H2O (pH 6.0)/-

II

N7 N7 N7 N7 N7 N7 N7 N7

HgII (2 mM) HgII (2 mM) ZnII (20 mM) ZnII (20 mM) ZnII (13.0 mM) ZnII (7.5 mM) CdII (5 eq.) PtII (15 mM)

MepdG (10 mM)/H2O (pH 7.5)/dGpMe (10 mM)/H2O (pH 7.5)/MepdG (10 mM)/H2O (pH 7.5)/dGpMe (10 mM)/H2O (pH 7.5)/d(ATGGGTACCCAT)2b (1.7 mM)/ H2O (pH 6)/d(ATGGGTACCCAT)2b (1.7 mM)/ H2O (pH 6)/r(GGACGAGUCC)2b/ H2O (pH 6.0)/d(TGGT) (15 mM)/ H2O (pH 6)/-

+0.69 – +0.88 +0.54

+0.8 −0.48 – −0.18 −0.65 0.0 +0.9 ∼−0.5

+2.4 +2.6 N.D. N.D.

C4

– – – – – – – −0.5

– – – –

HgII (0.7 M) HgII HgII (0.7 M) HgII (0.7 M)

guanosine (0.2 M)/DMSO/2′-deoxyguanosine/DMSO/inosine/DMSO/1-methylguanosine (0.2 M)/DMSO/-

+4.0 +3.90 +4.07 +5.69 +0.23 +3.9 +2.6 ∼+3

+2.7 +2.6 N.D. N.D.

C2

– – – – – – – –

N1 N1 N1 N1 N1 N1 N1 N1

HgII (0.5 eq.) HgII (0.7 M) HgII HgII (0.7 M) HgII (0.7 M) CH3HgII (1.0 eq.) CH3HgII (1.0 eq.) CH3HgII (1.0 eq.)

guanosine (isolated G-HgII-G)/DMSO/guanosine (0.2 M)/DMSO/Et3N (0.4 M) 2′-deoxyguanosine/DMSO/Et3N inosine/DMSO/Et3N 1-methylguanosine (0.2 M)/DMSO/Et3N (0.4 M) guanosine (isolated CH3HgII-G)/DMSO/inosine (isolated CH3HgII-I)/DMSO/5′-GMP (isolated CH3HgII-GMP)/H2O (pH 8.0)/-

N3 N3 N3 N3

HgII (0.5 eq.) CH3HgII (1.0 eq.) HgII (2.0 mM) HgII (2.0 mM)

Site

thymidine (isolated T-HgII-T)/DMSO/thymidine (isolated CH3HgII-T)/DMSO/d(GCGCTTTTGCGC)b (2 mM)/ H2O (pH 6.0)/d(GCGCTTTTGCGC)b (2 mM)/ H2O (pH 6.0)/-

Metal

C chemical shift perturbations upon metal-ion bindinga

13

Sample (M)/solvent/additive (M)

Table 16.1

446 Structural Studies on the MercuryII-Mediated T-T Base Pair

(2.0 mM)/H2O (2.0 mM)/H2O (2.0 mM)/H2O (2.0 mM)/H2O

(4.0 mM) (4.0 mM) (4.0 mM) (4.0 mM)

N1

N7 N7 N7 N7 N7 N7 N7 N7 N7 N7 N7 N7 N7 N7 N7

HgII (0.5 M) HgII (2 mM) HgII (2 mM) HgII (0.75 eq.) CdII (2.5 mM) CdII (12 mM) ZnII (0.7 eq.) ZnII (0.5 M) ZnII (20 mM) ZnII (20 mM) ZnII (4.9 mM) ZnII (4.9 mM) ZnII (12 mM) ZnII (1 eq.) ZnII (0.5 M)

guanosine (0.5 M)/DMSO MepdG (10 mM)/H2O (pH 7.5) dGpMe (10 mM)/H2O (pH 7.5) inosine/DMSO r(GGACGAGUCC)2b (0.5 mM)/ H2O (pH 6) r(CGGAGUUGGC) · r(GCCAAAGCCG)b (3 mM)/ H2O (pH 6.8) inosine/DMSO guanosine (0.5 M)/DMSO MepdG (10 mM)/H2O (pH 7.5) dGpMe (10 mM)/H2O (pH 7.5) d(GGTACCGGTACC)2b (0.29 mM)/H2O (pH 5.0) d(GGTACCGGTACC)2b (0.29 mM)/H2O (pH 5.0) r(CGGAGUUGGC) · r(GCCAAAGCCG)b (3 mM)/ H2O (pH 6.8) guanine (DFT calculation) adenosine (0.5 M)/DMSO

∼+1 −18.0 −11.9 ∼−22 ∼−0.5 – – ∼0 +2.7 – ∼0 ∼0 – – – – – – ∼0

−9.5 −1.0 +1.5 ∼−1 ∼0 – – ∼0 −0.6 – ∼0 ∼0 – – – – – – −1.6

∼+2.5 – – n.a. +3.1 – n.a. ∼+2 – – – – – – +1.4

+7.4 +1.2 ∼+10

∼+2

N.D. N.D.

−3.3 +2.4

+49.9 +15.3 +9.4

– – – –

NH2

∼+4

+30.3 +35.3 +29.9 +30.8

N3

∼+1

– – – –

N1

−1.2

−0.5 −0.7

∼+2

∼−20.5 −1.5 −0.5 −4.8 −19.6 −20.5 ∼−15 ∼−20 −8.4 −4.2 −4.0 −14.8 −20.0 −14.8 −7.5

n.a. n.a. n.a.

– – – –

N7

∼+4 N.D. N.D. ∼+2 +2.5 – ∼+1 ∼+3 +0.9 +1.0 <1 <1 – – ∼0

n.a. n.a. n.a.

∼+2.5

−3.0 −1.0

∼+3

– – – –

N9

62 65 65 63 75 78 63 62 65 65 74 74 78 79 62

62 64 62

62

69 69

64

83 83 83 83

Reference

a Chemical shift changes are listed in ppm. Positive and negative values represent lower-field shifts and higher-field shifts, respectively. MepdG: methylester of 5′-GMP; dGpMe: methylester of 3′-GMP; n.a.: not applicable; N.D.: not detected; –: not recorded. b For DNA/RNA oligomers, the chemical shift changes of the underlined residues are indicated.

N3 N3 N3

Hg (0.75 M) CH3HgII (0.2 eq.) ZnII (0.75 M)

cytosine (0.75 M)/DMSO 3′-CMP/H2O (pH 6.0) cytosine (0.75 M)/DMSO

II

HgII (0.5 M)

N1

N3 N3 N3 N3

adenosine (0.5 M)/DMSO

II

CH3HgII (1.0 eq.)

Hg HgII HgII HgII

Site

N6 N6

6) 6) 6) 6)

Metal

Hg (28 mM) HgII (28 mM)

(pH (pH (pH (pH

II

d(CGCGAATTCGCG) (4.8 mM)/H2O (pH 7) d(CGCGAATTCGCG) (4.8 mM)/H2O (pH 7)

b 2 b 2

5′-GMP (isolated CH3HgII-GMP)/H2O (pH 8.0)

d(CGCGTTGTCC) · d(GGACTTCGCG) d(CGCGTTGTCC) · d(GGACTTCGCG)b d(CGCGTTGTCC) · d(GGACTTCGCG)b d(CGCGTTGTCC) · d(GGACTTCGCG)b

b

N chemical shift perturbations upon metal-ionbindinga

15

Sample (M)/solvent

Table 16.2

NMR Spectral Studies 447

448

Structural Studies on the MercuryII-Mediated T-T Base Pair

NMR chemical shifts and signal widths (n1/2) of seven types of methylmercuryII– nucleobase complexes in DMSO solution.66 Among them, there were 199Hg NMR data for the methylmercuryII complex with uridine or thymine at N3 (type-V), and those with guanosine at N1. Next, in 1984, they reported NMR studies on a U-HgIIU pair with a C5-HgII-N3 linkage, and a U-HgII–6-thioguanosine pair with a C5HgII-S6 linkage67 In 1983, Buchanan and Bell reported a 199Hg NMR chemical shift of the inosine–HgII complex.63 Other sets of 199Hg NMR data of nucleobase complexes were reported by Lippert and coworkers.21–25,27 One is from the HgII–guanine derivative complexes,22 and another is from HgII,PtII bimetallic complexes with cytidine.21,27 From their studies, one-bond 199Hg–195Pt J-coupling (2783 Hz), five-bond 199 Hg–1H J-coupling (20 Hz for 199Hg-N4-C4-C5-C6-1H6),21 four-bond 199Hg–1H Jcoupling (78 Hz for 199Hg-N3-C4-C5-1H5)27 and three-bond 199Hg–1H J-coupling (107–222 Hz for 199Hg-C5-C6-1H6)23–25 were detected. These multinuclear NMR data provide important structural information for investigating more complicated systems.

16.4.2 Oligonucleotide–Metal Systems The first NMR data on HgII–DNA complexes were reported by Young and coworkers in 1982.68 In their report, they recorded 1H NMR spectra of poly-d(AT) in the presence of various concentrations of HgII, and obtained the following data: (i) as the concentration of HgII was increased, imino proton resonances were decreased and disappeared at a molar ratio of 0.25 for [HgII]/[per-nucleotide concentration]; (ii) average chemical shifts of nonexchangeable protons (H8 and H2 of adenine (A), and H6 and 5-CH3 protons of T) showed sigmoidal transitions against the molar ratio of [HgII]/[per-nucleotide concentration]; (iii) obtained data supported a strandslippage model with the formation of T-HgII-T base pairs. Studies on the HgII–oligonucleotide systems were then reported by Sletten and coworkers (Table 16.2).69–72 They studied two self-complementary DNA duplexes, d(CGCGAATTCGCG)2 and d(GCCGATATCGGC)2. It was found that HgII bound to AT-rich regions of both duplexes. In the first sequence, 1H and 15N NMR spectroscopic techniques were employed to elucidate the HgII binding mode in DNA duplexes. The most striking data are natural abundance 1H-15N HMQC spectra69,71 in which cross peak detection is extremely difficult because of the low sensitivity of 15 N nuclei. Sletten and coworkers overcame this issue by using a high concentration sample (4.8 mM duplex: d(CGCGAATTCGCG)2) with 1H-15N HMQC spectra using a two-bond 1H–15N J-coupling, and so made it possible to detect 15N resonances of N1 of cytidine, N7 and N9 of purine residues (adenosine and guanosine) and N1 and N3 of A.69,71 This technique has also been applied to other metal–DNA systems.73,74 From the data, it was found that very large lower-field shifts were observed at the N1 resonances of A5 and A6 residues (15.3 ppm and 9.4 ppm, respectively) (Table 16.2).69,71 It should be noted that Sletten and coworkers concluded that the observed lower-field shift would be attributed to HgII binding to amino nitrogen (N6) with proton–HgII exchange, after careful consideration. This is because for the

NMR Spectral Studies H N N R

II

N Hg N

N

449

O

H N O

N R

Scheme 16.3

coordination of N1 of A [N1(A)] to HgII, a higher-field shift is plausible for the 15N resonance of imine nitrogen (Table 16.2).62–64,75–79 It was also proposed that the bound HgII would be ligated with carbonyl oxygen (O4) of T [O4(T)] in the opposite strand.69–72 As a result, A-HgII-T base pairs were thought to be formed in the DNA duplex (Scheme 16.3). This structure was supported by HgII titration experiments with 1H NMR spectra of d(GCCGATATCGGC)2.70,72 From the 1H NMR data, imino proton resonances of thymine residues in A-HgII-T base pairs were observed after HgII binding was saturated.70,72 This means that imino protons of thymine residues were preserved after A-HgII-T base pair formation (Scheme 16.3). Thus, all NMR data are consistent with the proposed structure of the A-HgII-T base pair. Interestingly, this pairing arrangement is similar to the one proposed by Davidson’s group (Scheme 16.2),11 but the correct structure of the A-HgII-T base pair might be the one by Sletten’s group (Scheme 16.3). From these data, it was found that in the A,G,C,T mixed sequence, HgII can penetrate between a Watson– Crick A-T base pair without strand slippage or imino proton–HgII exchange processes. 16.4.3 T-HgII-T Base Pair In 1996, Kuklenyik and Marzilli reported a possible formation of T-HgII-T base pairs in DNA oligomers using 1H NMR spectroscopy.80 They performed HgII titration experiments with several DNA oligomers, namely, d(ATGGGTTCCCAT), d(GCGCTTTGCGC) and d(GCGCTTTTGCGC). As the concentration of HgII was increased, the imino proton resonances of underlined thymines decreased, and then disappeared around 1 molar equivalent to the T-T mismatches in the following context.80 For d(ATGGGTTCCCAT) and d(GCGCTTTGCGC), they seemed to form self-associated duplexes in the presence of HgII.80 In the case of d(GCGCTTTTGCGC) a hairpin structure was formed in the presence of HgII.80 However, on the addition of excess HgII, this hairpin molecule seemed to be transformed into a self-associated duplex gradually.80 However, the disappearance of the imino proton resonances suggested T-HgII-T base pair formation in the DNA duplexes. Thus, this NMR study provides the first NMR data for a possible formation of T-HgII-T base pairs in DNA oligomers with defined chain lengths and sequences. Similar HgII titration experiments were presented in 2006, and imino proton resonances of thymidine residues were seen to disappear upon the addition of approximately 1 molar equivalent of HgII to a T-T mismatch in a DNA duplex:

450

Structural Studies on the MercuryII-Mediated T-T Base Pair

d(CGCGTTGTCC) · d(GGACTTCGCG).81 For this DNA duplex, the sequences were designed specifically for the structure determination of the T-HgII-T base pair. On the addition of HgII, two successive T-HgII-T base pairs are formed in the middle of the duplex. It should also be mentioned that the sequence of the DNA duplex used in this experiment was made nonself-complementary, in order to avoid the loss of structural information. This is because if duplexes with self-complementary sequences are used, NOEs between symmetry-related protons around the dyad axis of the duplexes will be lost due to the averaging of NMR signals. By contrast, with the above nonself-complementary sequences, NOE cross peaks were not lost, and unambiguous resonance assignments based on NOEs were able to be achieved.82 During HgII titration experiments on the duplex, imino proton signals from II Hg -free duplex and HgII–duplex (1 : 1) complexes (transient complexes) were independently observed.81 It was found that association/dissociation processes are slow relative to the NMR timescale. Therefore, the disappearance of imino proton signals upon the addition of HgII supports the formation of the T-HgII-T base pair, as was the case in the studies by Kuklenyik and Marzilli.80 For nonexchangeable protons, such as the methyl protons of thymidine [CH3(T)], NMR signals from the HgII-free duplex, HgII-duplex (1 : 1) complexes and the HgII-duplex (2 : 1) complex were independently observed.82 More interestingly, 15N NMR spectra of the HgII complex of the same sequence was reported in 2007 (Table 16.2).83 In that study, 15N–15N J-coupling across HgII (2JNN) was observed within the T-HgII-T base pairs in the duplex, providing direct evidence of the formation of an 15N-HgII-15N covalent linkage (Scheme 16.4, Figure 16.4 and Table 16.3).83 At this stage, the chemical structure of the T-HgII-T base pair was definitely determined. Interestingly, a theoretical value of 2JNN for the T-HgII-T base pair was also reported by Bagno and Saielli in 2007; this showed a similar value to the experimental data (Table 16.3).84 For reference data, related metal-mediated 15 N–15N J-coupling values85,86 and trans-hydrogen bond 15N–15N J-coupling values87–89 are also listed in Table 16.3. To evaluate chemical shift perturbations upon HgII binding, chemical shifts of II Hg -free thymidine residues were determined using 1H-15N HSQC spectra (Figure 16.5).83 From the spectra, N3 resonances in the T-T mismatches (HgII-free form) were assigned, and the chemical shift differences between HgII-free and HgII-bound forms were determined (Table 16.2). It was found that approximately 30 ppm lowerfield shifts were observed for the N3 resonances of thymidines upon HgII binding (Table 16.2). This large chemical shift change is discussed later.

O 15

R

N

O II

N Hg O

2J

NN

N

15

O

Scheme 16.4

N R

NMR Spectral Studies

451

Figure 16.4 One-dimensional (1D) 15N NMR spectra of duplex-HgII (1 : 2) complexes: (a) duplex (1·3)-HgII complex; (b) duplex (2·3)-HgII complex; (c) duplex (2·4)-HgII complex. The inset of panel (c) shows the resolution-enhanced spectrum of the N3 resonance of T16 [N3(T16)]. The coupling constant of 15N-HgII-15N (2JNN) is shown in Hz as an absolute value. N3 resonances of thymidines are labelled with coloured circles. Each colour represents the position of the residue. 15N-frequency (0 ppm) is 81.07646745 MHz. (Reprinted with permission from J. Am. Chem. Soc., 2007, 129, 244–245. Copyright 2007 American Chemical Society.) Table 16.3 J-coupling valuesa Bond

2

JNN / Hz

Reference

15

2.4b 2.4b 1.7b +4.3 +4.3 +7 ∼ +8

83 83 84 85 86 87–89

N(T5)-HgII-15N(T16) N(T6)-HgII-15N(T15) 15 N(T)-HgII-15N(T) (DFT calculation) 15 N-RuII-15N (trans)c 15 N-FeII-15Nd 15 N(T)-H···15N(A)e 15

a

J-couplings are listed in Hz; Absolute values are listed; J-coupling from [RuCl(PPh3)(BPM)2]+Cl−, where BPM is bis(1-pyrazolyl)methane; d J-coupling from [Ph3PNPPh3]+ · [Fe4(NO)7S3]− e Trans-hydrogen bond J-coupling from N-H···N hydrogen bonds in Watson–Crick type A-T base pairs. b c

452

Structural Studies on the MercuryII-Mediated T-T Base Pair 150

duplex (1.3)

15 N

155

(ppm)

T5, T16

160 150

duplex (2.3)

15N

155

(ppm)

T6, T16

160 150

duplex (2.4)

11.0

10.5 1H (ppm)

10.0

15N

155

(ppm)

T6, T15

160 9.5

Figure 16.5 1H-15N HSQC spectra of duplexes (1·3), (2·3) and (2·4). By using these three spectra, we can assign each cross peak. For example, in the top and middle panels, the N-H cross peak of the T16 residue should be observed. Therefore, the cross peak at 10.2 ppm (1H), 154.4 ppm (15N) is the N-H cross peak of T16. Similarly, the cross peak at 10.3 ppm (1H), 153.9 ppm (15N) is that of T6. Finally, the cross peaks at 10.9 ppm (1H), 155.1 ppm (15N) and at 10.7 ppm (1H), 155.8 ppm (15N) are those of T5 and T15, respectively. (Reprinted with permission from J. Am. Chem. Soc., 2007, 129, 244–245. Copyright 2007 American Chemical Society.)

16.5 Relationship Between 15N Chemical Shifts and Chemical Bonds We investigated theoretical aspects of chemical shifts of N3 resonance at the HgII binding site to understand chemical reasons why the N3 resonances shifted toward such a low field upon HgII binding. From the definition of a chemical shift, the

15

N Chemical Shifts and Chemical Bonds 453

chemical shift of a target nucleus (dtarget) can be calculated using the following equations:90,91

δ target = σ ref − σ target

(16.1)

σ target = σ D + σ P

(16.2)

σD ∝ ρ

(16.3)

σ P ∝ 1 ∆E

(16.4)

where starget = shielding constant of the target nucleus, sref = shielding constant of the nucleus in the reference compound, r = electron density and ∆E = average excitation energy of electronic transitions. For details, please refer to references 90 and 91. In the case of 15N nuclei, their shielding constants include both s D and s P terms, which is a very different situation from those of 1H. More importantly, the s P term generally has a larger effect on s target than the s D term.90,91 Therefore, as a very rough approximation, the s15N is proportional to the reciprocal of ∆E:

σ 15N ≈ σ 15P N ∝ 1 ∆E

(16.5)

This average excitation energy (∆E) is highly correlated with the HOMO– LUMO transition or the allowed transitions near the HOMO–LUMO transition, since such transitions are the most frequently occurring transitions. This is the reason why the average excitation energy, ∆E, is highly correlated with the energy of the HOMO–LUMO or related excitations. For T-HgII-T base pair formation, the reaction goes though the pathway shown in Scheme 16.5. From the 15N chemical shifts of N3, HgII-bound N3 nuclei were resonated at a lower-field than proton-bound N3 nuclei. This means that s P of HgII-bound N3 nuclei [s P(N-HgII)] makes a larger contribution than s P of proton-bound N3 nuclei [s P(N-H)] (Figure 16.6).

σ P( N−Hg II ) > σ P( N−H )

(16.6)

Caution must be taken with the sign of s P, since its sign is opposite to the sign of s D. According to Equations (16.5) and (16.6), this phenomenon means that the excitation energy of the electron around the HgII-bound N3 nuclei [∆E(N-HgII)] would be smaller than the proton-bound N3 nuclei [∆E(N-H)] (Figure 16.6). O

R

N

O

H+

NH O

H+

R

N

O

HgII

N O

Scheme 16.5

HgII

R

N

N HgII O

454

Structural Studies on the MercuryII-Mediated T-T Base Pair

LUMO(+n) ∆E(N:-) O N

∆E(N-H)

O

∆E(N-HgII)

Stabilization by N-HgII bonding

Stabilization by N-H bonding

N

HOMO(-n)

N

HOMO(-n)

HOMO(-n)

O

O

N H

N

N

O

O

Figure 16.6 Theoretical explanation for the

O HgII

15

∆E ( N−Hg II ) < ∆E ( N−H )

N

N

HgII

O

N chemical shifts

(16.7)

This would be probably caused by a smaller binding energy for HgII to N3 than the proton-N3 binding energy (Figure 16.6). Interestingly, this interpretation is consistent with a reversible HgII–DNA interaction, since the activation energy (the energy for the HgII-N3 dissociation in this case) must be small enough for reversible transitions.

16.6 Applications of the T-HgII-T Base Pair Since the T-HgII-T base pair is a rather irregular base pair with extraordinary stability and HgII specificity, it can be applied to biochemical tools or in devices for nanotechnology.92–99 For example, the T-HgII-T base pair can be used as a nanowire made of a DNA molecule with T-HgII-T base pairs. Indeed, short DNA duplexes have been made of only T-HgII-T base pairs.81 Since DNA itself is a weak conductor, a metallated DNA duplex might be a good conductor. To shed light on this issue, Joseph and Schuster examined the hole-transport activity of a DNA molecule with T-HgII-T base pairs.92 From their studies, it was determined that a DNA molecule with T-HgII-T base pairs is also a hole transporter with similar activity to that of normal DNA molecules. However, it is also true that a DNA molecule with T-HgII-T base pairs is not as good a hole transporter. Theoretical calculations (DFT calculations) on the charge transfer through the DNA duplex with T-HgII-T base pairs were performed by Voityuk, in 2006.93 From his data, no obvious HgII-HgII interaction was found within HOMO or HOMO-n. Interestingly, this calculation result is consistent with the experimental data of Joseph and Schuste.92 It was, however, found that an HgII-HgII interaction was able to be seen in the LUMO, which suggested

Biological Relevance of the T-HgII-T Base Pairs

455

that a nanowire made of T-HgII-T base pairs might be a mediator for excess electron transfer.93 Utilizing the HgII specificity of the T-HgII-T base pair, DNA oligomer-based II Hg sensors were made by several groups. In 2004, an HgII sensor with fluorescent probes (a fluorophore at the 3′-end and a quencher at the 5′-end) was reported.94 In this sensor, the sensor–DNA molecule forms a hairpin structure upon HgII binding, which induces quenching of the fluorescence. Therefore, the concentration of HgII can be quantitated from the fluorescence intensity (the degree of quenching). As mentioned above, T-T mismatches are highly specific to HgII, and this HgII sensor specifically detected HgII in the presence or absence of various metal cations.94 In 2007, gold nanoparticles were utilized as an HgII sensor in a study that employed two kinds of gold nanoparticles modified with different DNA oligomers on their surfaces.95 The first particle is modified with d(AAAAAAAAAATAAAAAAAA AA), and the second with d(TTTTTTTTTTTTTTTTTTTTT). In the presence of HgII, a T-HgII-T base pair and Watson–Crick base pairs are formed between the DNA oligomers on the particles. This induces the aggregation of the gold nanoparticles and the colour of the particle-containing solution changes (colorimetric sensing).95 A third sensor was made from a catalytic DNA molecule (DNAzyme) and its substrate RNA molecule with fluorescent probes (a fluorophore at the 5′-end and a quencher at the 3′-end).96 In the presence of HgII, the DNAzyme strand takes an active structure and cleaves the substrate RNA, which leads to an increase in fluorescence intensity. The interesting point of the enzymatic sensor is that the HgII detection signal (fluorescence intensity) can be enlarged upon HgII sensing.96

16.7 Biological Relevance of the T-HgII-T Base Pairs NMR, FTIR and Raman studies have revealed modes of interaction between HgII and nucleobases. Among these structures, a A-HgII-T base pair with a N6(A)-HgIIO4(T) and a T-HgII-T base pair with a N3(T)-HgII-N3(T) can be formed under physiological conditions.69–72,81–83,94 The A-HgII-T base pair with the N6(A)-HgIIO4(T) linkage was found to exist in studies by Sletten’s group. This result indicates that, in the case of A,G,C,T-mixed DNA sequences, T-HgII-T base pairing with strand displacements is not necessarily a favourable process. Instead, penetration of HgII into Watson–Crick A-T base pairs can take place. Thus it is possible that reversible HgII binding to natural DNA molecules (E. coli DNA, calf thymus DNA, etc.) might involve such a mode of interaction. It has also been demonstrated that a T-HgII-T base pair with a N3(T)-HgII-N3(T) does exist, and its chemical structure was clearly determined in a DNA duplex.83 However, the proton–HgII exchange has not been observed in crystal structures of functional RNA molecules and DNA/RNA oligomers. Even in the cases with coordination bonds, those between metal cations and nucleic acids are rarely observed. Therefore, the observation of the T-HgII-T base pair in the context of the DNA duplex suggests that metal cation-binding with nucleic acids though a proton–HgII exchange can occur if the conditions are appropriate for such proton–HgII exchanges.

456

Structural Studies on the MercuryII-Mediated T-T Base Pair

•••ATATATAT••• n•HgII 2n•H+

T

•••TATATATA•••

T

A A

T T

A A

T T

A A

T T

HgII

Figure 16.7 A possible mechanism for strand slippage

: HgII poly-T (or T-rich) sequence

n•

2n•H+

T-HgII-T base-pairs

poly-dA (or A-rich) sequence poly-dA (or A-rich) loop

Figure 16.8 A possible mechanism of deletion mutation by HgII

This interaction mode can rationalize the strand-slippage model for polyd(AT)-HgII complex with a T-HgII-T base pair. This is because only in the case of the dA-dT alternating polymer [poly-d(AT)], does strand slippage by one base lead to a continuous duplex with A-A mismatch pairs and T-HgII-T base pairs (Figure 16.7). It has also been suggested that T-HgII-T base pairing might be related to deletion mutations when human cells are exposed to a high concentration of HgII (Figure 16.8).100 As a possible mechanism for this mutation, poly-d(A)-poly-d(T) sequences might form T-HgII-T base pairs within the poly-d(T) strand to give a hairpin loop. The resulting looped-out region would be skipped by DNA polymerase during replication of the genome. Therefore, from a toxicological point of view, the HgII– DNA interaction would be related to biological phenomena.28,100,101 Here, we discuss the thermodynamic stability of the T-HgII-T base pair. Upon T-HgII-T base pairing, the Tm of the DNA duplex, d(CGCGTTGTCC) · d(GGAC TTCGCG), is considerably higher than that of the full-matched DNA duplex, d(C GCGTTGTCC) · d(GGACAACGCG) (YT unpublished data). It was found that the T-HgII-T base pair has a similar (or higher) thermodynamic stability, relative to Watson–Crick base pairs. Therefore, this thermodynamic stability might be a driving force for strand slippage/displacement and loop-out. Next, we would like to note the kinetics of T-HgII-T base pairing. From NMR studies, the HgII association/dissociation processes in DNA duplexes have been shown generally to be slow relative to an NMR timescale.80–83 These observations are consistent with the kinetic data from Williams and Crothers, who reported stopped-flow experiments with UV spectra.102 Therefore, once a T-HgII-T base pair is formed, the pairs are not only thermodynamically, but also kinetically, stable, suggesting that it is difficult to dissociate T-HgII-T base pairs once they are accidentally formed in cells. For this reason, biological organisms possess a defense system against toxic metal cations (HgII, CdII, PbII and so on), such as metallothioneins (metal-ion-

References

457

chelating proteins). Furthermore, in the case of eukaryotic cells, genomic DNA is surrounded by a nuclear membrane and is shielded from HgII attack. Therefore, effects of HgII on the genetic mutations might be evident in prokaryotes. In addition, intrinsic genetic effects of HgII could be studied by using PCR as the simplest model replication system.

16.8 Concluding Remarks The structure of the T-HgII-T base pair, once one of the mysteries of HgII–DNA interaction modes in DNA duplexes, was determined from 15N NMR spectra. Especially, 15N–15N J-coupling across HgII (2JNN: 15N-HgII-15N) provided definite evidence of the N-HgII-N covalent linkage in the T-HgII-T base pair. Interestingly, its structure was fully consistent with the structural model proposed by Katz in 1963. It is astonishing that even in the early stages of these studies, the correct structure of the THgII-T base pair was suggested from limited data. It is also interesting to note that the lower-field shifts of 15N NMR chemical shifts upon HgII binding are consistent with the reversibility of the N-HgII bond formation. Therefore, the N-HgII bond might be classified into a relatively weak category among covalent bonds. However, T-HgII-T base pairs with N-HgII bonds are sufficiently stable to form a duplex that is rather more stable than the Watson–Crick A-T base pair in some sequences. Furthermore, its association/dissociation kinetics are slow relative to an NMR timescale. Such characteristics of the T-HgII-T base pair would promote irregular DNA structures. However, little is known about the weak interatomic interactions within the T-HgII-T base pair and interactions between neighbouring T-HgII-T base pairs. These residual issues need attention as the next targets in this field.

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29. Gruenwedel, D.W.; Effect of Hg(II) on the spectroscopic properties of poly[d(A-T)·d(AT)] and Poly[d(A)·(T)] and their constituent subunits (deoxyadenosine and thymidine monomers and dimers); Biophys. Chem., 1994, 52, 115–123. 30. Gruenwedel, D.W.; Cruikshank, M.K.; Smith, G.M.; Effect of Hg(II) on d(GCGCATATGCGC)2 conformation – UV absorption and circular-dichroism studies; J. Inorg. Biochem., 1993, 52, 251–261. 31. Gruenwedel, D.W.; Effect of Hg(II) on the spectroscopic properties of DNA bases – circular-dichroism of deoxyadenosine and thymidine monomers and dimers; J. Inorg. Biochem., 1994, 56, 201–212. 32. Gruenwedel, D.W.; The Mercury(II) and high-salt-induced conformational B-Z transitions of poly[d(G-M(5)C) · d(G-M(5)C)] as studied by non-polarized (ultraviolet) and circularly-polarized (CD) ultraviolet spectroscopy; Eur. J. Biochem., 1994, 219, 491–496. 33. Mansy, S.; Wood, T.E.; Sprowles, J.C.; Tobias, R.S.; Heavy metal-nucleotide interactions. Binding of methylmercury(II) to pyrimidine nucleosides and nucleotides. Studies by raman difference spectroscopy; J. Am. Chem. Soc., 1974, 96, 1762–1770. 34. Mansy, S.; Tobias, R.S.; Heavy metal-nucleotide interactions. II. Binding of methylmercury(II) to purine nucleosides and nucleotides studied by raman difference spectroscopy; J. Am. Chem. Soc., 1974, 96, 6874–6885. 35. Mansy, S.; Tobias, R.S.; Metal ion catalysis of proton exchange between purine nucleosides or nucleotides and solvent water; J. Chem. Soc. Chem. Commun., 1974, 957– 958. 36. Mansy, S.; Frick, J.P.; Tobias, R.S.; Heavy metal-nucleotide interactions III. The participation of amino-groups in the binding of methylmercury(II) to cytidine and adenosine 5′-phosphate in aqueous-solution: Studies by raman difference spectrophotometry; Biochim. Biophys. Acta, 1975, 378, 319–332. 37. Mansy, S.; Tobias, R.S.; Heavy metal-nucleotide reactions 4. Nature of reaction between mercury(II) and uridine or thymidine – vibrational spectroscopic studies on binding to N(3), C(4) = O, and C(5) of uracil base; Inorg. Chem., 1975, 14, 287–291. 38. Mansy, S.;Tobias, R.S.; Heavy metal nucleoside interactions. Binding of methylmercury(II) to inosine and catalysis of isotopic-exchange of C-8 hydrogen studied by 1H nuclear magnetic resonance and raman difference spectrophotometry; Biochem., 1975, 14, 2952–2961. 39. Chrisman, R.W.; Mansy, S., Peresie, H.J., Ranade, A., Berg, T.A., Tobias, R.S.; Heavy metal-nucleotide interactions 9. Raman difference spectroscopic studies on binding of CH3Hg(II) to 1-methylthymine, thymidine-5′-monophosphate, DNA models and native DNA; Bioinorg. Chem., 1977, 7, 245–266. 40. Canty, A.J.; Tobias, R.S.; Heavy-metal-nucleoside interactions. 13. Synthesis and spectroscopic study of organomercury derivatives of guanosine and thyidine; Inorganic Chemistry, 1979, 18, 413–417. 41. Savoie, R.; Jutier, J.-J.; Prizant, L.; Beauchamp, A.L.; Raman and infrared spectra of methylmercury complexes of adenine; Spectrochim. Acta. A, 1982, 38, 561–568. 42. Morzyk-Ociepa, B.; Michalska, D.; Vibrational spectra of 1-methylthyminate complexes with mercury(II) and potassium and ab initio calculations of the 1-MeT anion; J. Mol. Struct., 2001, 598, 133–144. 43. Morzyk-Ociepa, B., Michalska, D.; Vibrational spectra of 1-methyluracilate complex with silver(I) and theoretical studies of the 1-MeU anion; Spectrochim. Acta. A, 2003, 59, 1247–1254. 44. Morzyk-Ociepa, B., Nowak, M.J., Michalska, D.; Vibrational spectra of 1-methylthymine: matrix isolation, solid state and theoretical studies; Spectrochim. Acta. A, 2004, 60, 2113–2123. 45. Szczepaniak, K.; Szczesniak, M.M.; Person, W.B.; Raman and infrared spectra of thymine. A matrix isolation and DFT study; J. Phys. Chem. A, 2000, 104, 3852–3863. 46. Wang, S.W.; Li, N.C.; Proton magnetic resonance studies of metal complexes of imidazole purine and pyrimidine derivatives; J. Am. Chem. Soc., 1966, 88, 4592–4597.

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47. Wang, S.W.; Li, N.C.; Proton magnetic resonance studies of self-association and metal complexation of nucleosides in dimethyl sulfoxide; J. Am. Chem. Soc., 1968, 90, 5069–5074. 48. Kang, L.S.; Li, N.C.; Nuclear magnetic resonance studies of mercury(II) interaction with nucleosides in dimethyl sulfoxide; J. Am. Chem. Soc., 1970, 92, 4823–4827. 49. Tewari, K.C.; Lee, J.; Li, N.C.; Zinc and mercury(II) interaction with cytidine and glycylglycine; Trans. Faraday Soc., 1970, 66, 2069–4597. 50. Shimokawa, S.; Fukui, H.; Sohma, J.; Hotta, K.; Proton magnetic-resonance studies of metal complexation of nucleosides in dimethyl sulfoxide; J. Am. Chem. Soc., 1973, 95, 1777–1782. 51. Chang, C.-H.; Marzilli, L.G.; Nucleoside complexing. A charge reversed chelate compound between guanosine and chloride ion; J. Am. Chem. Soc., 1974, 96, 3656–3657. 52. Yokono, T.; Shimokawa, S.; Sohma, J.; Nuclear magnetic-resonance studies on effects of ions (anions and cations) to nucleosides (cytidine and guanosine); J. Am. Chem. Soc., 1975, 97, 3827–3829. 53. Marzilli, L.G.; Stewart, R.C.; Van Vuuren, C.P.; de Castro, B.; Caradonna, J.P.; Metal ion binding to cytidine in solution. Compelling raman and carbon-13 nuclear magnetic resonance spectral evidence for coordination to the exocyclic oxygen at position 2; J. Am. Chem. Soc., 1978, 100, 3967–3968. 54. Marzilli, L.G.; de Castro, B.; Caradonna, J.P.; Stewart, R.C.; Van Vuuren, C.P.; Nucleoside complexing – raman and C-13 NMR spectroscopic study of the binding of hard and soft metal species; J. Am. Chem. Soc., 1980, 102, 916–924. 55. Marzilli, L.G.; Metal complexes of nucleic acid derivatives and nucleotides: binding sites and structures; Adv. Inorg. Biochem., 1981, 3, 47–85. 56. Marzilli, L.G.; de Castro, B.; Solorzano, C.; Nucleoside complexing: A 13C NMR spectroscopic study of binding of metal-ions to guanosine and related nucleosides in solution – evidence for 06 binding under basic conditions; J. Am. Chem. Soc., 1982, 104, 461–466. 57. Buncel, E.; Norris, A.R.; Racz, W.J.; Taylor, S.E.; Metal ion-biomolecule interactions. Synthesis, spectroscopic, and magnetic resonance investigations of methylmercury(II) complexes of the nucleosides guanosine and inosine; Inorg. Chem., 1981, 20, 98–103. 58. Buncel, E.; Boone, C.; Joly, H.; Kumar, R.; Norris, A.R.; Metal ion-biomolecule interactions. Part 12. 1H and 13C NMR evidence for the preferred reaction of thymidine over guanosine in exchange and competition reactions with mercury(II) and methylmercury(II); J. Inorg. Biochem., 1985, 25, 61–73. 59. Buncel, E.; Boone, C.; Joly, H.; Metal-ion biomolecule interactions. Part 13. NMR evidence for the formation of the mixed-ligand thymidine-mercury-guanosine complex – a model for a putative Hg(II) interstrand cross-linking structure of DNA; Inorg. Chim. Acta., 1986, 125, 167–172. 60. Buncel, E.; Norris, A.R.; Racz, W.J.; Taylor, S.E.; Metal ion–C(8) binding in purine nucleosides. Ready formation of carbon-bound inosine and guanosine complexes of methylmercury(II); J. Chem. Soc. Chem. Commun., 1979, 562–563. 61. Taylor, S.E.; Buncel, E.; Norris, A.R.; Metal ion-biomolecule interactions. II. Methylmercuration of the deprotonated amino groups in adenine, guanine, and cytosine derivatives, and its relationship to amino group acidity; J. Inorg. Biochem., 1981, 15, 131–141. 62. Buchanan, G.W.; Stothers, J.B.; Diamagnetic metal-ion nucleoside interactions in solution as studied by 15N nuclear magnetic resonance; Can. J. Chem., 1982, 60, 787–791. 63. Buchanan, G.W.; Bell, M.J.; 15N and 199Hg magnetic resonance spectral investigation of inosine protonation and diamagnetic metal-ion complexation; Can. J. Chem., 1983, 61, 2445–2448. 64. Buchanan, G.W.; Bell, M.J.; Mercuration effects on nucleoside chemical shifts – 15N and 13 C NMR studies of methylmercury(II) ion addition to cytidine-3′-monophosphate and guanosine-5′-monophosphate; Magn. Reson. Chem., 1986, 24, 493–497. 65. Polak, M.; Plavec, J.; Multinuclear NMR study of the conformational changes in MepdG and dGpMe upon interaction with Mg2+, Zn2+ and Hg2+ ions reveals the strengthening of the anomeric effect by soft M2+ ions; Eur. J. Inorg. Chem., 1999, 547–555.

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66. Norris, A.R.; Kumar, R.; 199Hg NMR correlations in methylmercury(II) complexes of nucleic-acid constituents and their analogs; Inorg. Chim. Acta., 1984, 93, L63–L65. 67. Norris, A.R.; Kumar, R.; Metal-ion biomolecule interactions. Part IX. Metal-ion C-5 binding in a pyrimidine nucleoside – ready formation of C-5-Hg-N and C5-Hg-S Bonds; Inorg. Chim. Acta, 1984, 93, 33–35. 68. Young, P.R.; Nandi, U.S.; Kallenbach, N.R.; Binding of mercury(II) to poly(dA-dT) studied by proton nuclear magnetic resonance; Biochem., 1982, 21, 62–66. 69. Frøystein, N.Å.; Sletten, E.; Interaction of mercury(II) with the DNA dodecamer CGCGAATTCGCG – A 1H and 15N NMR study; J. Am. Chem. Soc., 1994, 116, 3240–3250. 70. Steinkopf, S.; Nerdal, W.; Kolstad, A.; Sletten, E.; Sequence-selective interaction between mercury(II) ions and the DNA dodecamer [d(GCCGATATCGGC)](2) studied by H-1 NMR spectroscopy; Acta Chim. Scand., 1996, 50, 775–782. 71. Sletten, E.; Frøystein, N.Å.; NMR studies of oligonucleotide-metal ion interactions. In Metal Ions in Biological Systems, Vol. 32, Sigel, A. and Sigel, H., Eds., 1996, 397–418. 72. Sletten, E.; Nerdal, W.; Interaction of mercury with nucleic acids and their components. In Metal Ions in Biological Systems, Vol. 34, Sigel, A. and Sigel, H., Eds., 1997, 479–501. 73. Vinje, J.; Parkinson, J.A.; Sadler, P.J.; Brown, T.; Sletten, E.; Sequence-selective metalation of double-helical oligodeoxyribonucleotides with PtII, MnII, and ZnIIiIons; Chem. Eur. J., 2003, 9, 1620–1630. 74. Vinje, J.; Sletten, E.; Internal versus terminal metalation of double-helical oligodeoxyribonucleotides; Chem. Eur. J., 2006, 12, 676–688. 75. Tanaka, Y.; Kojima, C.; Morita, E.H.; Kasai, Y.; Yamasaki, K.; Ono, A.; Kainosho, M.; Taira, K.; Identification of the metal ion binding site on an RNA motif from hammerhead ribozymes using 15N NMR spectroscopy; J. Am. Chem. Soc., 2002, 124, 4595–4601. 76. Tanaka, Y.; Kasai, Y.; Mochizuki, S.; Wakisaka, A.; Morita, E.H.; Kojima, C.; Toyozawa, A.; Kondo, Y.; Taki, M.; Takagi, Y.; Inoue, A.; Yamasaki, K.; Taira, K.; Nature of the chemical bond formed with the structural metal ion at the A9/G10.1 motif derived from hammerhead ribozymes; J. Am. Chem. Soc., 2004, 126, 744–752. 77. Tanaka, Y.; Taira, K.; Detection of RNA nucleobase metalation by NMR spectroscopy; J. Chem. Soc. Chem. Commun. 2005, 2069–2079. 78. Wang, G.; Gaffney, B.L.; Jones, R.A.; Differential binding of Mg2+, Zn2+, and Cd2+ at two sites in a hammerhead ribozyme motif, determined by 15N NMR; J. Am. Chem. Soc., 2004, 126, 8908–8909. 79. Sychrovsky, V.; Sponer, J.; Hobza, P.; Theoretical calculation of the NMR spin-spin coupling constants and the NMR shifts allow distinguishability between the specific direct and the water-mediated binding of a divalent metal cation to guanine; J. Am. Chem. Soc., 2004, 126, 663–672. 80. Kuklenyik, Z.; Marzilli, L.G.; Mercury(II) site-selective binding to a DNA hairpin. Relationship of sequence-dependent intra- and interstrand cross-linking to the hairpinduplex conformational transition; Inorg. Chem., 1996, 35, 5654–5662. 81. Miyake, Y.; Togashi, H.; Tashiro, M.; Yamaguchi, H.; Oda, S.; Kudo, M.; Tanaka, Y.; Kondo, Y.; Sawa, R.; Fujimoto, T.; Machinami, T.; Ono, A.; Mercury(II)-mediated formation of thymine-HgII-thymine base pairs in DNA duplexes; J. Am. Chem. Soc., 2006, 128, 2172–2173. 82. Tanaka, Y.; Yamaguchi, H.; Oda, S.; Nomura, M.; Kojima, C.; Kondo, Y.; Ono, A.; NMR spectroscopic study of a DNA duplex with mercury-mediated T-T base pairs; Nucleosides Nucleotides Nucleic Acids, 2006, 25, 613–624. 83. Tanaka, Y.; Oda, S.; Yamaguchi, H.; Kondo, Y.; Kojima, C.; Ono, A.; 15N-15N J-coupling across HgII: direct observation of HgII-mediated T-T base pairs in a DNA duplex, J. Am. Chem. Soc., 2007, 129, 244–245. 84. Bagno, A.; Saielli, G.; Metal-mediated J-coupling in DNA base pairs: Relativistic DFT predictions; J. Am. Chem. Soc., 2007, 129, 11360–11361. 85. Otting, G. ; Messerle, B.A.; Soler, L.P.; 1H-detected multinuclear NMR experiments for the measurement of small heteronuclear coupling constants in transition metal complexes; J. Am. Chem. Soc., 1997, 119, 5425–5434.

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86. Butler, A.R.; Glidewell, C.; Hyde, A.R.; Homonuclear spin spin coupling through iron in simple inorganic-ions; Polyhedron, 1984, 3, 1165–1167. 87. Dingley, A.J.; Grzesiek, S.; Direct observation of hydrogen bonds in nucleic acid base pairs by internucleotide 2JNN couplings; J. Am. Chem. Soc., 1998, 120, 8293–8297. 88. Pervushin, K.; Ono, A.; Fernández, C.; Szyperski, T.; Kainosho, M.; Wüthrich, K.; NMR scaler couplings across Watson-Crick base pair hydrogen bonds in DNA observed by transverse relaxation optimized spectroscopy; Proc. Natl. Acad. Sci. USA, 1998, 95, 14147–14151. 89. Barfield, M.; Dingley, A.J.; Feigon, J.; Grzesiek, S.; A DFT study of the interresidue dependencies of scalar J-coupling and magnetic shielding in the hydrogen-bonding regions of a DNA triplex; J. Am. Chem. Soc., 2001, 123, 4014–4022. 90. Levy, G.C.; Lichiter, R.L.; Nitrogen-15 Nuclear Magnetic Resonance Spectroscopy, John Wiley & Sons, Inc., New York, 1979. 91. de Dios, A.C.; Ab initio calculations of the NMR chemical shift; Prog. Nucl. Magn. Reson. Spectr., 1996, 29, 229–278. 92. Joseph, J.; Schuster, G.B.; Long-distance radical cation hopping in DNA: The Effect Of Thymine-Hg(II)-thymine base pairs; Org. Lett., 2007, 9, 1843–1846. 93. Voityuk, A.A.J.; Electronic coupling mediated by stacked [thymine-Hg-thymine] base pairs; Phys. Chem. B, 2006, 110, 21010–21013. 94. Ono, A.; Togashi, H.; Highly selective oligonucleotide-based sensor for mercury(II) in aqueous solutions; Angw. Chem. Int. Ed., 2004, 43, 4300–4302. 95. Lee, J.-S.; Han, M.S.; Mirkin, C.A.; Colorimetric detection of mercuric ion (Hg2+) in aqueous media using DNA-functionalized gold nanoparticles; Angew. Chem. Int. Ed., 2007, 46, 4093–4096. 96. Liu, J.; Lu, Y.; Rational design of ‘turn-on’ allosteric DNAzyme catalytic beacons for aqueous mercury ions with ultrahigh sensitivity and selectivity; Angew. Chem. Int. Ed., 2007, 46, 7587–7590. 97. Torigoe H, Kawahashi K, Takamori A, Ono, A.; Novel strategy for single nucleotide polymorphism (SNP) genotyping by heteroduplex analysis: Specific stabilization of TT mismatch base pair by mercury (II) cation and CC mismatch base pair by silver (I) cation; Nucleosides Nucleotides Nucleic Acids, 2005, 24, 915–917. 98. Miyake Y, Ono, A.; Fluorescent sensor for redox environment: a redox controlled molecular device based on the reversible mercury mediated folded structure formation of oligothymidylate; Tetrahedron Lett.; 2005, 46, 2441–2443. 99. Tanaka, K.; Clever, G.H.; Takezawa, Y.; Yamada, Y.; Kaul, C.; Shionoya, M.; Carell, T.; Programmable self-assembly of metal ions inside artificial DNA duplexes; Nature Nanotech., 2006, 1, 190–194. 100. Ariza, M.E.; Williams, M.V.; Lead and mercury mutagenesis: Type of mutation dependent upon metal concentration; J. Biochem. Mol. Toxicol., 1999, 13, 107–112. 101. Müller, J.; Sigel, R.K.O.; Lippert, B.; Heavy metal mutagenicity: insights from bioinorganic model chemistry; J. Inorg. Biochem., 2000, 79, 261–265. 102. Williams, M.N.; Crothers, D.M.; Binding kinetics of mercury(II) to polyribonucleotides; Biochem., 1975, 14, 1944–1951. 103. Jia, X.; Zon, G.; Marzilli, L.G.; Multinuclear NMR investigation of Zn2+ binding to a dodecamer oligodeoxyribonucleotide – insights from 13C NMR spectroscopy; Inorg. Chem., 1991, 30, 228–239. 104. Mukundan, S. Jr., Xu, Y., Zon, G. and Marzilli, L.G.; Heteronuclear 13C-1H NMR investigation of the effects on an oligodeoxyribonucleotide of intrastrand cross-linking by a Pt anticancer drug – a large shift of C3′ accompanies an S-conformational to Nconformational change; J. Am. Chem. Soc., 1991, 133, 3021–3027. 105. Tanaka, Y.; Morita, E.H.; Hayashi, H.; Kasai, Y.; Tanaka, T.; Taira, K.; Well-conserved tandem G · A pairs and the flanking C · G pair in hammerhead ribozymes are sufficient for capture of structurally and catalytically important metal ions; J. Am. Chem. Soc., 2000, 122, 11303–11310.

17 Chromium-Induced DNA Damage and Repair Laura G. Little and Kent D. Sugden

17.1 Introduction 17.1.1 Chromium as a Carcinogen and Current Exposure Limits Chromium was first used as a commercial chemical in the early 1800s.1 Since that time, chromium has found an ever increasing use in a wide range of industrial processes. These include commercial leather tanning, pigment production, corrosion inhibition, chrome plating, wood preservation, photoengraving, lithography and fireworks production.2 It was not until the late 19th century, however, that the hexavalent oxidation state of chromium – Cr(VI) or chromate-containing chemicals were noted as a potential human health hazard. Scottish chrome pigment workers began reporting a large number of nasal tumours presumably caused by the inhalation of chromate dust.3 By the 1930s, epidemiological studies in Germany had noted a link between Cr(VI) exposure and the development of lung cancers.4 These findings were later confirmed by studies in the United States,5–7 United Kingdom,8–11 Germany,12 Italy13 and Japan.14 Compounds containing Cr(VI) are now listed as Class 1 human carcinogens by the International Agency for Research on Cancer (IARC)15 and as a Group A airborne human carcinogen by the US EPA due to their presence in a number of occupational settings.16–18 Safe exposure limits for Cr(VI) compounds are still being adjusted as new information becomes available. In 1992, the drinking water standard for total chromium was lowered to 0.1 ppm, due to the emergence of Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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new information about amounts of chromium being released into the environment.19 OSHA has recently settled on a more stringent new standard for occupational exposure to hexavalent chromium. The new standard has lowered the permissible exposures from 52 µg m−3 to 5 µg m−3 of air as an 8 hour time-weighted average.20 These recent changes in Cr(VI) exposure limits highlight the increasing concern for the health and safety of occupationally and environmentally exposed human populations. 17.1.2 Physical Properties of Chromium that Confer Carcinogenicity Like most transition metals, chromium can exist in a number of oxidation states. Known oxidation states for this metal range from −II to +VI.21 The most common oxidation states are III and VI but Cr(III) is naturally more abundant than Cr(VI) due to its higher stability in a reducing environment.2 Oxidative processing of the more abundant Cr(III) ores to industrially useful compounds of Cr(VI) have been extensive in many of the industries listed above. Waste from those processes artificially increases the amount of Cr(VI) released into the environment, which is of increasing concern because of the carcinogenic nature of these Cr(VI) compounds with respect to the more benign Cr(III) form. However, Cr(VI) is not the oxidation state of chromium that directly interacts with nucleic acids to initiate cancer formation.22 Instead, it is the intracellularly reduced oxidation states of Cr(V), Cr(IV) and Cr(III) that have been observed to cause damage to DNA.22 Hexavalent chromium compounds are much more carcinogenic than trivalent chromium compounds because Cr(VI) and not Cr(III) is able to rapidly cross the cell membrane through relatively nonselective anion channels23 that normally regulate the uptake of phosphate and sulfate ions. At physiological pH, Cr(VI) exists as a chromate oxyanion, which is of a similar size and has the same tetrahedral geometry as phosphate and sulfate ions under those same conditions (Figure 17.1).3 The octahedral Cr(III) ions O HO -O

Cr

O

pH > 7

+6 O

-O -O

pH < 7

HCrO4-

O

Chromate L

L

+6

CrO42-

Hydrogen Chromate

L

Cr

+3

Cr

L L

L

CrL6 Chromium(III) Figure 17.1 The major forms of physiologically relevant chromium species

Introduction

465

are also able to cross the cell membrane, but rely solely on passive diffusion and are therefore taken up approximately three orders of magnitude more slowly than Cr(VI) ions.24 Once in the cell, Cr(VI) is quickly reduced by a variety of intracellular reductants including ascorbic acid and reduced thiols, such as cysteine and the ubiquitous tripeptide glutathione.25 Ascorbic acid is the most kinetically favoured of these reductants26 and is thought to be the major intracellular reductant of Cr(VI). However, the exact contribution of the different reductants towards intracellular chromate reduction and genotoxicity is not known. Irrespective of the exact nature of the reductant, the ensuing rapid intracellular reduction of Cr(VI) to Cr(V), Cr(IV) and finally to Cr(III) creates a concentration gradient across the cell membrane that favours the continual uptake of Cr(VI) from the extracellular matrix. To adequately understand the mechanism of how chromium interacts with DNA it is first necessary to define the DNA reactive intermediates that are formed during the intracellular reduction process of Cr(VI). 17.1.3 Reactive Intermediates Formed During Reduction of Chromate The determination of a mechanistic pathway of cancer formation by Cr(VI) has been complicated by the wide variety of reactive species formed during the intracellular metabolism of this metal. These pathways are further complicated by the fact that many of these reactive intermediates may give rise to similar DNA-damage products, making it a challenge to associate specific reactants with the final premutagenic DNA lesion. As stated above, there are three intracellular chromate reductants that have been widely studied and are considered to be the primary players in chromate metabolism and the ensuing DNA damage. These are ascorbate, glutathione and, to a lesser degree based on low physiological concentration, hydrogen peroxide. Reduction of chromate by all three of these compounds forms both radical species and high valent metal species. While most research has focused on the radical-mediated pathways, it is becoming clear that a direct or metal-mediated pathway may play as great or greater role in DNA damage induced by chromate. Cr(VI) Reactions with Ascorbate The in vitro reaction of Cr(VI) with ascorbate has been extensively studied. Ascorbate is kinetically one of the best physiological reductants of chromate and was found to be the major reductant in rat liver, lung and kidney ultrafiltrates.27 Cr(VI) reduction by ascorbate has been found to form a number of different radical species including the ascorbyl radical and the carbon dioxide radical cation (Figure 17.2; I and II respectively) as well as high valent ascorbate-ligated Cr(V) and Cr(IV) species (Figure 17.3A). The relative yield of each of these reactive species is dependent upon ascorbate to Cr(VI) ratios.28,29 However, since Cr(VI) uptake is essentially steady-state from the extracellular milieu, and ascorbate concentrations in human cells are variable depending on dietary status, defining a Cr(VI):ascorbate ratio and the likely reactive intermediates that will be formed is unpredictable at best. Gross DNA-damage studies using plasmid DNA however, have implicated the high-valent metal species generically shown in Figure 17.3A as the likely reactive intermediates

466

Cr-Induced DNA Damage and Repair Carbon-centred radicals from ascorbate O

O

+ O

CO2

OH

O OH

I

II

Oxygen-centred radicals from hydrogen peroxide +

1

O2

+

OH

O2

IV

III

V

Sulfur-centred radical from glutathione H N

O

O O

NH

O

S

H3N

O

VI

O

Figure 17.2 Compilation of reductant-based radical species observed upon reduction of chromate

O

O

+5

O

Cr

A)

+4

O O

O

O O

O

O S

B)

N

O

Cr

O

O

+5 Cr

+4

S

N

N

Cr N

S

S O

O O

C)

O

O Cr

O

O

+5 O O

Figure 17.3 Putative high-valent intermediates of chromium arising during the intracellular reduction of chromate by: (A) ascorbic acid, ( B) glutathione and (C) hydrogen peroxide

Introduction

467

responsible for lesion formation.30–32 Perhaps more interesting is current research suggesting that ascorbate, classically thought of as an antioxidant, may serve as a pro-oxidant with regard to intracellular metabolism of Cr(VI).33 There is also recent evidence that ascorbate-ligated high-valent chromium species may have multiple roles in both oxidative DNA damage and in the formation of Cr-DNA adducts, as detailed in later sections of this review. Cr(VI) Reactions with Glutathione Glutathione, a tripeptide of g -glutamylcysteinylglycine, is likely to be another major player in the intracellular reduction of Cr(VI). Reduction of Cr(VI) in the presence of glutathione is significantly slower than that observed with ascorbate, but glutathione is generally present in cells at considerably higher (millimolar) levels than ascorbate.34 A number of reactive intermediates have been associated with the Cr(VI) metabolism process by glutathione.35–37 Among these are the glutathionyl radical (Figure 17.2; VI), which may react with molecular oxygen to produce the superoxide anion38 (Figure 17.2; V). Similar to the reactions observed with ascorbate metabolism of Cr(VI), glutathione metabolism of this metal also forms high-valent Cr(V) species (Figure 17.3B) that have been observed by electron spin resonance spectroscopy, ESR.35,36 Once again, the ultimate species that may be responsible for DNA damage during the glutathione-dependent metabolism process is unknown. Much like ascorbate, glutathione’s normal role of maintaining cellular redox homeostasis may be altered to one of a pro-oxidant with regard to Cr(VI) metabolism. Cr(VI) Reactions with Hydrogen Peroxide The reactions of Cr(VI) with hydrogen peroxide have been extensively studied, although this may be due more to the ability of this reaction to induce significant DNA modifications than with any cellular reality. Hydrogen peroxide is highly reactive with metal species and as such is kept at very low levels in the cell, which would seem to preclude any significant metabolic reduction of Cr(VI) by this species.39,40 However, a case could be made for reactions of this nature in the mitochondria where a steady-state level of hydrogen peroxide is produced during respiration accounting for up to 5% of total cellular oxygen consumption.41 In general, the reaction of Cr(VI) with hydrogen peroxide generates the classical reactive oxygen species, ROS, that have been observed with the Fenton metals of iron and copper. These include singlet oxygen, the hydroxyl radical and the superoxide anion (Figure 17.2; III, IV and V respectively).42,43 A high-valent Cr(V) tetraperoxochromate species (Figure 17.3C) has also been observed by ESR that is highly reactive with DNA.44 This species is only formed in extremely alkaline solutions and is probably not physiologically relevant. The DNA reactivity of ROS species have been extensively reviewed elsewhere and will not be discussed to any length in this review. However, it should be noted that many of the same DNA lesions formed by these classical ROS are now being found to arise from a direct or metal-mediated type of oxidation reaction with DNA.

468

Cr-Induced DNA Damage and Repair

17.1.4 General Types of DNA Damage Associated with Chromate A wide variety of genetic damage that may initiate cancer formation has been observed following Cr(VI) exposure. This wide array of genetic damage has confounded the chromium research community and there is still little general consensus on the mechanism(s) of DNA damage by Cr(VI), even though it is one of the most extensively studied carcinogenic metals. This damage includes DNA base and sugar oxidation, Cr-DNA adducts, DNA–protein crosslinks, DNA–DNA crosslinks and DNA strand scission.45 The downstream effects of these lesions, if not outright toxicity, can be lesion accumulation, mutation and apoptosis resistance. Cells that do not undergo apoptosis due to the accumulation of chromium-induced genetic damage will develop gene mutations,46 chromosomal aberrations47,48 and sister chromatid exchanges.47–49 This genetic damage caused by hexavalent chromium exposure is of considerable concern for both industrial chrome workers and the general population. Even with improved industrial practices, some workers are still exposed to as much as several hundred µg of Cr(VI) m−3 of air.47 The general public is also at risk due to environmental chromium pollution from a legacy of 150 years of often illconceived landfill practices, as well as pollution stemming from the mining, smelting and wood treatment industries. There are 719 active superfund sites in the United States with chromium listed as at least one of the major contaminants present.50 Automobile emissions and cigarette smoke are another source of nonoccupational exposure to Cr(VI).47 Understanding the mechanisms and types of chromiuminduced genotoxicity is essential for combating the effects of occupational and environmental exposure to Cr(VI) compounds. The remainder of this review focuses on specific types of DNA damage that Cr(VI) compounds are known to form, the current knowledge of how endogenous repair processes play a role in the removal of these adducts and, the induction of downstream cellular effects from these DNAdamaging processes.

17.2 Nucleobase Oxidation 17.2.1 Cr(VI)-Induced Damage at DNA Nucleobases Once Cr(VI) has been taken up by a cell, it is rapidly reduced.25 The highly reactive, metastable intermediate oxidation states of Cr(V) and Cr(IV) are created as Cr(VI) is reduced to the more stable Cr(III).51–53 This reduction scenario sets up two major pathways that can lead to oxidation of the nucleobases. One pathway involves the formation of radical species as detailed above, but with primary emphasis placed on reactive oxygen species (ROS). Through Fenton-like reactions with H2O2, both Cr(III)53 and Cr(VI)54,55 have been shown to be capable of producing hydroxyl radicals that can attack the electron-rich aromatic rings of the nucleobases. The ROS pathway is favourable in the presence of high concentrations of H2O2 (0.5 mM– 27 mM),53–55 which have been used for many in vitro studies. However as stated previously, under normal physiological conditions the concentration of H2O2 in a

Nucleobase Oxidation

469

cell is much lower than 1 mM,56 indicating that oxidation of nucleic acids by ROS is likely to play a small role in the overall nucleobase oxidative damage that has been observed in cells exposed to Cr(VI). A second pathway that is thought to account for the majority of nucleobase oxidation induced by Cr(VI) exposure involves the direct abstraction of an electron from nucleic acids by the Cr(V) and/or Cr(IV) intermediate species formed during Cr(VI) reduction.57 The vast majority of nucleobase oxidation observed during the Cr(VI) reduction process occurs at guanine residues. Guanine is preferentially oxidized by ROS and by high-valent chromium intermediates because of its lower one-electron reduction potential relative to that of the other three bases (Table 17.1).58–60 The reduction potential becomes even lower for 5′ guanine residues when a run of consecutive guanine residues is present in a DNA sequence (Table 17.1).58–60 The most recognized lesion formed by a single electron abstraction mechanism from a guanine residue is 7,8-dihydro-8oxo-2′-deoxyguanosine (8-oxoG) (Figure 17.4). In fact, 8-oxoG is often viewed as the primary genetic marker of oxidative stress23 and if not repaired is mutagenic, as 8-oxoG can mispair with adenine during DNA replication leading to G:C→T:A transversion mutations.61 Table 17.1 Reduction potentials for nucleosides and nucleotide sequences58–60 Nucleosidea

E (V vs NHE)

DNA Sequence

E (V vs NHE)

Guanine (G) Adenine (A) Cytosine (C) Thymine (T) 8-oxoG

1.29 1.42 1.6 1.7 0.7

GGG GG GA GC GT

0.64 0.82 1.00 1.15 1.16

a

The reduction as listed is for the radical cation of the corresponding nucleoside or nucleotide.

O N N R

NH N

O

O Crn

NH2

Crn-1

N N R

OH

NH N

NH2

H

N

HO

N R

NH N

NH2

- e-

Guanine

- H+ O H N

NH

O N R

N

NH2

8-oxoG

Figure 17.4 Sequential single electron oxidation mechanism of a guanine residue that gives rise to the premutagenic 8-oxoG lesion

470

Cr-Induced DNA Damage and Repair O

N N R

O NH

N Guanine

LxCrn NH2

O L Crn x

N

NH

O N R

O

LxCrn-2 O

N

NH2

H N N R

NH N

NH2

8-oxoG

Figure 17.5 Two-electron oxidation mechanism that gives rise to 8-oxoG

High-valent metal species such as those observed to arise from Cr(VI) reduction (Figure 17.3) have the potential to generate the same 8-oxoG lesion, but through a different mechanism than that of the single-electron oxidation shown in Figure 17.4. This mechanism involves the transfer of the axial oxo atom (a twoelectron oxidation mechanism) of the Cr(V) or Cr(IV) complex to the substrate through a discrete innersphere interaction (Figure 17.5). While this mechanism has yet to be shown to form 8-oxoG from guanine, it has been shown to occur for a number of other substrates.62,63 Validation of this mechanism is possible using isotopically labelled oxygen in the oxo-atom-containing chromium complex, but is problematic due to the rapid exchange of the oxo-atom with the surrounding water.62 However, a similar mechanism for 8-oxoG formation has been observed with highvalent ruthenium-oxo complexes.64 For years 8-oxoG was considered the terminal (and most relevant) oxidation product of guanine in DNA. However, the reduction potential of 8-oxoG is considerably lower than that of the parent guanine residue (Table 17.1) making it a prime target for further oxidative events. While it may seem unlikely for two discrete oxidative events to occur at the same DNA base (without some sort of a cage effect) recent studies have shown the formation of further oxidized products of 8-oxoG both in vitro65 and in cellular systems.66 A mechanism referred to as charge transfer has been proposed to explain this effect. When an oxidative event occurs at one base in a DNA sequence it can be translated through the DNA to an electron sink – a nucleoside with a lower reduction potential than the neighbouring nucleosides.67 The stacked bases in B-form DNA create an array of overlapping π-orbitals that facilitate charge transfer through the DNA sequence.68 8-OxoG lesions can act as a sink for charge transfer because of their lowered reduction potential relative to that of the four undamaged nucleosides. When an 8-oxoG lesion is present within approximately 10–20 base pairs (presumably from an initial oxidative insult and greatly dependent on the DNA sequence)68 of the site of a secondary electron abstraction from a nucleobase, the resultant charge can be transferred to an 8-oxoG residue, causing a further oxidative event at that lesion. The low reduction potential of 8-oxoG combined with its ability to act as a sink for charge transfer in a DNA sequence suggests that 8-oxoG is not likely to be the end product of an oxidative attack on the nucleobases. One of the further oxidized lesions that has been shown to form both in vitro65 and in cellular systems66 is spiroiminodihydantoin (Sp). Sp lesions can be generated by a single electron abstraction from 8-oxoG (Figure 17.6),65 which can be achieved by both Cr(V) and Cr(IV)

Nucleobase Oxidation O H N

- e-

NH

O N

N R

NH2

8-oxoG

N

H N

- H+

NH

O N R

O

O H N

471

- e-

N

O N R

NH2

N

NH2

H2O H

O O HN O

NH N R

N H

acyl shift NH

Spiroiminodihydantoin, Sp

H N

O

O N

O N R

N

NH2

5-OH-8-oxoG

Figure 17.6 Single-electron oxidation mechanism of 8-oxoG to give the Sp lesion65

intermediates during reduction by ascorbate. In this mechanism, the initial oneelectron abstraction forms an 8-oxoG radical cation intermediate that is a substrate for nucleophilic addition of water to form the 5-hydroxy-8-oxoG intermediate that undergoes rearrangement via an acyl shift to give Sp (Figure 17.6). It should be noted that while the overall reaction is a two-electron oxidation, the mechanism invokes two discrete one-electron abstraction processes. Recently, an alternative mechanism has been proposed for the formation of the further oxidized product Sp from 8-oxoG.69 This mechanism is similar to that for the formation of 8-oxoG from guanine through the oxo atom transfer mechanism proposed in Figure 17.5. In this mechanism, an inner-sphere oxidation complex is formed with the 8-oxoG lesion, resulting in transfer of the oxo atom from the Cr(V) species to the C5 of 8-oxoG (Figure 17.7).69 This mechanism was confirmed using isotopic labelling, although it is still unclear whether it is viable in cellular systems since it would require a direct interaction between the chromium complex and the 8-oxoG lesion. The formation of Sp helps to explain the high level of mutations, and the varying types of mutations, observed in Cr(VI) treated systems, that cannot be accounted for by the formation of 8-oxoG lesions alone. Sp, like 8-oxoG, can produce G:C→T:A transversion mutations, but at a much higher level than 8-oxoG. In addition to the G:C→T:A transversion mutations, Sp can also produce G:C→C:G transversion mutations and polymerase arrest both in vitro and in cellular systems.70–72 There are potentially a wide variety of nucleobase lesions that have the ability to be formed from multiple chromium-induced oxidation events at guanine in DNA. 8-OxoG and Sp are the only two that have been observed in cellular systems and as such are the main focus in this part of the review. A lesion similar to Sp, guanidinohydantoin (Gh), has been shown to form with Cr(V) treatment in vitro from the 8-oxoG nucleoside,73,74 but has yet to be identified as arising in cellular systems. Figure 17.8 shows a more comprehensive picture of the wide variety of nucleobase

Cr-Induced DNA Damage and Repair

472

H N

X NH

O N R

O

O

O

N

RN

Cr X

+5 X

X

-2e-

NH

N

O O

NH2

N H

8-oxoG

O

O

X

Cr X 5+

N

O

H N N R

HO-

HN O

N N R

N H

H

acyl shift

HO

O

H O N

NH2

NH

N R

H2O

O

H N

O N

NH

O N R

OH-

NH2

NH2

N

nucleophilic attack ring opening

OO

NH

N

NH2

OH

Spiroiminodihydantoin, Sp Figure 17.7 Formation of Sp from 8-oxoG via a two-electron, oxo atom transfer mechanism69 O

O O

H N

HN

NH

O N R

NH2

N 8-oxoG

N R

O

N H

NH2

Spiroiminodihydantoin, (Sp)

O

HN

N

O

HN

NH

O

NH

O N R

N H

NH2

N R

N

NH2

Guanidinohydantoin, (Gh) Oxidized Guanidinohydantoin, (Ghox) O

O N

RHN

O

NH2 NH2

NH2 N

RHN

Oxazolone (Oz)

Imidazalone (Iz) O

H N

O

N

O

O NH

RN O Parabanic Acid (Pa)

HO

RHN

O

Oxaluric Acid (Oa)

Figure 17.8 Oxidative nucleobase lesions considered possible to arise from Cr(VI) exposure. Only 8-oxoG and Sp have thus far been observed in cellular systems

Nucleobase Oxidation

473

lesions that may be induced by Cr(VI) exposure. It should be noted that, with the exception of Sp and Gh, none of these other further oxidized lesions have yet been identified as arising from Cr(VI) treatment of DNA in vitro or in cellular systems. They have, however, been identified as arising in vitro in a number of other oxidizing systems. 17.2.2 Repair of Nucleobase Damage Repair of oxidized nucleobases is necessary to prevent the perpetuation of mutations in the genetic code. Oxidized DNA lesions such as 8-oxoG and Sp are typically repaired via the base excision repair (BER) pathway. There are several glycosylases that have been shown to repair the oxidized nucleobase lesions induced by Cr(VI) exposure. Table 17.2 provides a list of those glycosylases and some of the lesions that they are known to recognize. OGG1 is a mammalian glycosylase which is a homologue of the bacterial BER enzyme MutM and functions to repair 8-oxoG lesions.75 OGG1-deficient mice have been shown to accumulate 8-oxoG lesions over their wild-type counterparts.76 Another bacterial BER enzyme, endonuclease VIII (Nei), has been recently characterized77 and shown to repair a number of further oxidized DNA lesions including Sp.78 Nei deficient Escherichia coli have been shown to accumulate Sp lesions, but not 8-oxoG lesions66 similar to that reported in OGG1deficient mice indicating that the Nei repair enzyme is directed at the repair of these secondary oxidation products. Recently, three mammalian homologues of the Nei gene were identified and labelled Neil1, Neil2 and Neil3 (Nei-like).79,80 The NEIL1 and NEIL2 proteins were initially found to recognize cytosine-derived lesions80 and 8-oxoG lesions present in DNA bubble structures generated during transcription or replication.81 Recently NEIL1 and NEIL2 have been shown to efficiently recognize the oxidized lesions Sp and Gh, but have very poor, to no, affinity for 8-oxoG in normal B-form DNA.82 NEIL1 was shown to cleave Sp and Gh in both singleand double-stranded oligonucleotides, while NEIL2 showed cleavage of Gh in both substrates, but could not cleave Sp in double-stranded oligonucleotides.82

Table 17.2 BER glycosylases known to repair Cr(VI) induced nucleobase lesions. Some of these glycosylases are known to repair additional lesions not listed here but those additional lesions have not yet been associated with Cr(VI) induced genetic damage Prokaryotic BER glycosylase

Known nucleobase substrates (for prokaryotic BER only)

Mammalian BER homologuesa

MutM (Fpg)

8oxoG:C, Sp:C, Gh:C, 8oxoG:G, Sp:G, Gh:G 8oxoG:A, G:A Sp:A, Gh:A, 8oxoG:C, Sp:C, Gh: C, 8oxoG:G, Sp:G, Gh:G

OGG1

MutY Nei

MYH NEIL1, NEIL2

a The mammalian BER homologues generally do not have as wide a range of nucleobase substrates as that of the prokaryotic BER enzymes shown.

474

Cr-Induced DNA Damage and Repair

Repair of isolated nucleobase lesions by BER glycosylases involves a baseflipping mechanism. A damaged guanine residue will be located by a glycosylase scanning the DNA. It will then be flipped out from the interior regions of the DNA strand and into the active site pocket of the attached glycosylase.83 This base-flipping mechanism allows the glycosylase to gain access to the glycosidic bond of the nucleobase lesion. Once the glycosidic bond has been cleaved, the base is released and the glycosylase forms a Schiff base at the abasic site.83 This Schiff base will remain in place until it is displaced by other proteins needed to further process the abasic site. These proteins include apurinic/apyrimidinic endonuclease 1 (APE1) which cleaves the phosphate backbone just to the 5′ side of the abasic site, DNA polymerases, which then fill in the gap in the DNA polymer, and finally DNA ligase, which repairs the nick in the phosphate backbone. Many BER glycosylases harbour intrinsic AP lyase activity, which would allow them to further process the abasic site. This activity is very weak in most glycosylases and they do not complete the belimination, but instead this cleavage step is typically carried out by APE1. NEIL1 and NEIL2 however, have strong lyase activity and are thought to complete b/delimination at the abasic site.84 Recently, NEIL1 and NEIL2 have shown to have considerably different roles with regard to replication associated repair (RAR). NEIL1, but not NEIL2, shows strong cell cycle dependence with the S-phase.85 NEIL1, but not NEIL2, has also shown enhanced activity in the presence of proliferating cell nuclear antigen (PCNA), a processivity factor for eukaryotic polymerase d. The NEIL1/PCNA-stimulated activity suggests that NEIL1 may play an active role in oxidized base lesion removal during replication.86 Recent modelling and dynamic studies on NEIL1 have shed light on why this BER enzyme is specific for the nonplanar, further oxidized lesions, such as Sp and other modified pyrimidines.80,81 These studies have shown that the binding pocket is somewhat flexible and is more accommodating of the pyrimidine-like further oxidized products, such as Sp and has complementary hydrogen-bond donor–acceptor properties.87 In contrast, the planar 8-oxoG lesion binds only shallowly into the pocket, has few hydrogen-bond donor–acceptor pairs, and has a solvent-exposed six-membered ring.87 Taken together, the recent data on the formation of further oxidized guanine lesions in DNA and the identification of specific repair mechanisms suggest that these lesions may be important in the genotoxicity of a variety of carcinogens, including Cr(VI).

17.3 Sugar Oxidation 17.3.1 Cr(VI)-Induced Damage at Deoxyribose Highly reactive Cr(V) and Cr(IV) species are capable of directly abstracting electrons from the conjugated nucleobases, as discussed above. Similarly, these chromium species can also oxidize the deoxyribose sugar by direct abstraction of hydrogen atoms. Studies have shown that chromium preferentially attacks the C4′

Sugar Oxidation

475

and C1′ hydrogens.57,88 This preference stems from the accessibility of the C4′ which, along with the C5′ hydrogens, project into the minor groove and are the most solvent accessible of the deoxyribose hydrogens in normal B-form DNA.86 Cr(V) is considered to preferentially attack the C4′hydrogen and not the C5′ hydrogen of deoxyribose because it is easier to remove a hydrogen from a tertiary rather than a secondary carbon despite the fact that the C5′ hydrogens are slightly more solvent accessible.88 While the tertiary C1′ hydrogen is also oriented towards the minor groove, it is not as readily accessible for oxidation by chromium, as H1′ abstraction requires that an oxidant bind to the minor groove and then orient itself directly towards the H1′ position.89 In the presence of hydrogen peroxide, (1 mM) abstraction of the C1′ hydrogen has been observed.90 Reactions with hydrogen peroxide are considered of little significance in the overall scheme of chromium-induced genetic damage, since physiological concentrations of hydrogen peroxide are much lower than 1 mM.56 However, a model metastable Cr(V) complex with the bis-tris buffer designated Cr(V)-BT has shown the formation of the deoxyribose oxidation product 5-methylene-2-furanone (5-MF) that would indicate a classical C1′ hydrogen abstraction mechanism (Figure 17.9).57 It is proposed that the correct orientation for hydrogen abstraction at C1′ by this Cr(V) complex is generated through interaction with the phosphate backbone of DNA.57 This Cr(V) complex also showed a smaller amount of C4′ oxidation product. A different model Cr(V) complex, bis(2-ethyl-2-hydroxybutyrato)oxochromat e(V) or Cr(V)-ehba (which is proposed to resemble the Cr(V)-ascorbate complexes formed during ascorbate-induced Cr(VI) reduction; Figure 17.3A), has been shown to abstract the C4′ hydrogen of deoxyribose. This C4′ hydrogen abstraction mechanism

OR O

OR O-

P

O

O H

O

H P OR

H H

O O

B

O-

Cr(V)-BT

B O-

P

+

O

B

O H

H H1'

H O

H

O

H

O P

O

O2

O-

5-methylene-2-furanone (5-MF)

OR + OR O

P

O-

OH free 3'-phosphate termini

Figure 17.9 Abstraction of the C1′ hydrogen by Cr(V)-BT leads to formation of the free base, the 5-methylene-2-furanone sugar product and frank strand scission57

476

Cr-Induced DNA Damage and Repair OR

O

P

OR O-

O

O H

O

H4' H P OR

O-

Cr(V)-ehba

P

O-

O H

H

O O

P

O2

H

H

H

B

O

H H

O O

B

B

H

H

H O

O-

Base propenal

OR

+ OR O

P

O-

O O

O

glycolic acid termini

Figure 17.10 Abstraction of the C4′ hydrogen by Cr(V)-ehba leads to formation of a base propenals and frank strand scission88,91

was assigned based on the formation of deoxyribose oxidation products of base propenals, glycolic acid (Figure 17.10) and, like the C1′ mechanism, gives rise to frank DNA strand breaks57,88,89,91 Also, much like the Cr(V)-BT model complex, Cr(V)-ehba has shown the ability to interact with phosphates to position the oxidizing metal towards the site of oxidation (the C4′ hydrogen atom of deoxyribose).92 Single-strand breaks, such as those initiated by abstraction of a C4′ or C1′ hydrogen by Cr(V) and/or Cr(IV) species, are one of the most common genetic lesions reported following exposure to Cr(VI).45 Single-strand breaks have been shown to cause base deletions,93 as well as gross chromosomal damage94 including sister chromatid exchanges47–49 and micronuclei formation.45 The gross chromosomal damage that has been observed in cells exposed to Cr(VI) has led to the labelling of hexavalent chromium compounds as clastogenic in addition to their classification as carcinogens. Another lethal downstream effect of single-strand breaks are doublestrand breaks created when a replication fork encounters a single-strand break and collapses.96 Based on the literature of Cr(VI)-induced single-strand breaks, it is apparent that rapid repair of these breaks is necessary to avoid genotoxicity. 17.3.2 Repair of Deoxyribose Oxidation Products Single-strand breaks occur frequently, even when a cell has not been exposed to a genotoxin and are in fact thought to be the most common type of genetic lesion.95

Chromium–DNA Binding

477

Cells have developed a repair system for single-strand breaks that employs the same proteins whether the damage is generated endogenously or exogenously.95 There are four steps involved in repair of single-strand breaks: detection, end processing, gap filling and ligation. Single-strand breaks are initially detected by poly (ADPribose) polymerases 1 and 2, which bind to the breakage site and recruit endprocessing proteins.91 Breaks induced by high-valent chromium species have a 3′ sugar fragment that can be removed by APE1 with the help of X-ray repair crosscomplementing protein 1 (XRCC1).93,95 Gap filling is accomplished by insertion of a single nucleotide followed by DNA ligase sealing the nick in the sugar backbone.

17.4 Chromium–DNA Binding 17.4.1 Formation of Chromium Adducts with DNA Chromium, whether it be in the +V, +IV or +III oxidation state, can be attracted to the negatively charged DNA–phosphate backbone. Chromium ions may associate themselves with DNA through initial electrostatic interactions with the phosphate backbone that can lead to the formation of a metal–ligand complex with the phosphate oxygens.92 However, the metal–ligand complex formed is generally transient in nature, as the electron lone pair necessary for coordination is shared between the two oxygens attached to the phosphate (Figure 17.11a). This type of chromium– DNA adduct has shown to be facile to exchange and nonmutagenic in cellular systems.97 As such, it is likely of little importance with regard to the overall mechanism of DNA damage by chromium that gives rise to carcinogenesis. Formation of covalent bonds between chromium and the DNA nucleobases is a more controversial proposal. A number of studies have suggested that chromium may interact with guanine in DNA at the N7 in much the same manner as that seen with cis-platinum.98 However, the lone pair of electrons at the N7 of guanine are delocalized into the purine ring and as such, coordination at this site would not be favoured for chromium. The unique acid/base properties, square planar geometry and Jahn–Teller distortions associated with cis-platinum that allow binding at the N7 of guanine in DNA99 do not exist with chromium. Despite DNA being a poor ligand, some studies have shown that a portion of reduced chromium appears to associate in a covalent manner with DNA. This binding appears only when the reduced chromium is formed during the reduction with Cr(VI) and not through interaction with the final, stable reduced chromium oxidation state, Cr(III). This suggests that the intermediate Cr(V) and/or Cr(IV) oxidation states of chromium may play a role in this binding mechanism. The DNA binding interaction with chromium has been proposed to be through the formation of ternary adducts, (Figures 17.11b,d).100 These ternary adducts involve bidentate chelation between the DNA phosphate backbone and the N7 group of a guanine residue. One study has shown that following treatment of DNA with Cr(VI) and a reductant, a large amount of chromium was initially associated with the DNA. However, a high salt wash removed all but 20% of the initial associated chromium. What chromium remained

Cr-Induced DNA Damage and Repair

478

O L L

R

L O

Cr L

N

P

L

NH

O N

O

O

O

L

N

O

O

a

NH

L NH2

N

O

H H

N

Cr

P

NH2

N

O

L

L

R

H

H

R

H

H H

R

H

H

H

b

H L R L O

P

L Cr

O L

O

O

H N

O N

O O H c

Figure 17.11 adducts100,102

H

R

N

L

R

NH

H

NH2

P

Cr

H

O

N

NH

L

O

N

O

NH2

N

O

H H

O

L

H

H

H

H

R

H

H

d

Proposed structures for covalent Cr(III)–DNA bound binary and ternary

associated with the DNA was chelatable with EDTA indicating that it was likely covalently bound.101,102 It should be noted, however, that no detailed structure has yet shown the veracity of these adducts. The ability of chromium to covalently bind to DNA would allow for the formation of chromium-mediated DNA–DNA and protein–DNA crosslinks that have been proposed to arise in Cr(VI)-treated cellular systems. This will be discussed in more detailed in subsequent sections. 17.4.2 MMR Repair and Toxicity Associated with Chromium–DNA Adducts Recent studies have indicated that the small ternary chromium–DNA adducts proposed in Figures 17.11b and d are recognized and repaired by the mismatch repair (MMR) system.103 Inactivating any of the four major MMR proteins (MSH6, MSH2, MLH1, PMS2) from human or mouse cells creates a Cr(VI) resistant phenotype.103,104 Cells proficient in MMR should undergo apoptosis following exposure to low doses of Cr(VI), as MMR plays a role in both repair and controlled cellular toxicity.103 MMR is proposed to recognize DNA bending caused by Cr–DNA adducts105 and then cleave a segment of DNA – up to 1000 base pairs – surrounding these adducts.106–108 This repair process is now thought to act as a signal for the activation of cell cycle arrest and eventually apoptosis by creating single-strand breaks,93 which are manifested into double-strand breaks following a round of replication.107 Because a round of replication is required before the double-strand breaks will be manifested, MMR-induced double-strand breaks appear most frequently in the G2 phase of the cell cycle.102,107 The ability of MMR to trigger G2 cell cycle arrest as well as

Chromium–DNA Binding

479

apoptosis is enhanced by supplementation of cell cultures with ascorbate.104,109 As mentioned earlier, ascorbate is thought to be the major intracellular reductant of Cr(VI) creating high-valent, highly reactive chromium intermediates. These studies further the hypothesis that ascorbate, normally viewed as an antioxidant, may act as a pro-oxidant in cells treated with Cr(VI). One specific characteristic of lung cancers caused by Cr(VI) – unlike other lung carcinogens – indicates that cells carrying MMR deficiencies will be prone to becoming cancerous. This characteristic is called microsatellite instability110 and is associated with a complete loss of MMR activity.111,112 Another unique characteristic of Cr(VI)-induced cancers is that lung tumours from Cr(VI) cancer subjects retain a wild-type p53 tumour suppressor gene.113 Therefore, constant exposure to Cr(VI) compounds may not be necessary for tumour formation, since one would thus expect the p53 gene to accumulate a large number of chromium-induced mutations. Cells lacking MMR show resistance to an initial pulse of Cr(VI). This would lead to accumulation of a small number of lesions and adducts, and then the lack of an MMR system would allow those cells to continue to generate spontaneous mutations without exposure to additional chromium. A model has been developed by Salnikow and Zhitkovich to explain how MMR deficient cells may actually be selected following exposure to hexavalent chromium compounds (Figure 17.12).113 Cr(VI) Exposure

Formation of Cr-DNA Adducts

MMR Proficient Cells

MMR Deficient Cells

G2 Cell Cycle Arrest

Apoptosis Resistance

Apoptosis

Cell Replication

Tumour Formation

Figure 17.12 Selection scheme for developing a pool of MMR-deficient cells that may explain the induction of a cancerous phenotype from chromate exposure113

480

Cr-Induced DNA Damage and Repair

Those cells that are MMR deficient will be apoptosis resistant following Cr(VI) exposure. While the normal MMR proficient cells undergo apoptosis, the now initiated MMR-deficient cells will continue to grow and divide creating a population of cells that harbour a host of genetic mutations. To date, these studies are the most complete that show a correlation between chromium adduct formation, the role of DNA repair and the downstream cellular effects that can lead to cancer formation.

17.5 Cr(VI)-Induced DNA Crosslinks 17.5.1 Formation of Crosslinks As mentioned earlier, another class of lesions that can form following exposure of DNA to high-valent chromium species are crosslinks. Crosslinking can occur between DNA and a protein, between two adjacent sites on the same DNA strand (intrastrand crosslink) or between neighbouring DNA strands (interstrand crosslink). Small amounts of chromium can covalently bind to DNA through interactions with the phosphate backbone116,117 and form ternary adducts as described above.100,102 Chromium(III)-mediated ternary adducts may also involve a linkage between the phosphate backbone and protein, or with adjacent or opposite N7 groups of guanine residues. These lesions would be the chromium-mediated protein and DNA crosslinks. Since the DNA phosphate backbone is a fairly weak ligand, crosslink formation involving covalently bound chromium atoms cannot fully explain the extensive formation of crosslinks observed in many chromium-treated systems. These type of coordinative adducts are the least well defined and, once again, no structural characterization have been carried out on these systems. An alternative proposal to explain the formation of protein and DNA crosslinks is an oxidative pathway. Formation of single-strand breaks due to deoxyribose oxidation57,88 and accumulation of 8-oxoG and Sp due to guanine oxidation82 following Cr(VI) exposure demonstrate that chromium can be a powerful DNA oxidant. Oxidation of the guanine residue, as well as the 8-oxoG residue, forms a radical cation, as depicted in Figures 17.4 and 17.6. The radical cation presents an excellent site for nucleophilic attack and, although water is not a strong nucleophile, the vast amounts of water present surrounding the DNA favours the formation of lesions such as 8-oxoG and Sp. A number of much better nucleophiles are also present around the DNA, but are substantially less abundant than water. These include amino acids, such as histidine and cysteine, either as small peptides or within complete proteins, as well as the phosphate backbone of neighbouring DNA strands. When these strong nucleophiles are in close proximity to a chromium-generated radical cation, they may out-compete water and a DNA adduct with that nucleophile will be formed rather than an oxidized base.69 Therefore, DNA–DNA and protein–DNA crosslinks could be formed without the discrete coordination of a chromium atom, but as a result of a single-electron oxidation induced by Cr(V) or Cr(IV) ions (Figure 17.13). While no structural evidence for this type of reaction in

Cr(VI)-Induced DNA Crosslinks 481 H N O N

O NH

N

N

dR Guanine

O

O

Cr(V)/Cr(IV)

H N

NH

O NH2

N

O

N

H N

Cr(V)/Cr(IV)

NH

O

NH2

dR

NH2

N

dR

8-oxoG

Protein or other nucleophile

R2

C

dR

N HN

NH2

Spiroiminodihydantoin, Sp

N

O

N

O H2 > 7 pH

O

HN CH CH2

R1 H N

H N

N

NH

dR

N O

O

O N

N H

NH2

Guanidinohydantoin, Gh

N

O N

H pH 2 O < 7

N

NH2

dR

DNA-ProteinCrosslink

Figure 17.13 Putative oxidative crosslinking mechanism showing the formation of a DNA– protein crosslink from the guanine oxidation pathway

DNA has yet been shown, the validity of this reaction with chromium has recently been demonstrated by Cr(V)-induced adduction of a lysine to an oxidized 8-oxoG nucleoside.69 17.5.2 NER and Recombination Repair of Crosslinks Regardless of how the crosslinks are formed, they greatly alter the normal conformation of B-form DNA. Such bulky lesions can prove to be lethal for the cell as they lea,d not only to premutagenic deletions, but also to replication and transcriptional blockage.45 Nucleotide excision repair (NER) and recombination repair are required to restore the DNA template to its undamaged form. Many protein crosslinks recruit NER mechanisms because they only form a minor distortion of the DNA phosphate backbone.117,118 This is because a large proportion of the protein crosslinks do not involve a complete protein, just individual amino acids or short peptide chains,119,120 creating a relatively compact lesion. These types of crosslinks create enough distortion to recruit NER repair proteins,45 but are not bulky enough to require recombination repair. There are five steps involved in NER: lesion recognition, unwinding of the damaged region, incision, excision and synthesis/ligation.121 High-valent chromiuminduced DNA–protein crosslinks are initially identified by Xeroderma pigmentosum group A (XPA)122 and replication protein A (RPA),123,124 which interact with one another and with the DNA lesion to enhance the specificity of lesion recognition.125 The XPA protein will then recruit Xeroderma pigmentosum group B (XPB) and

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Xeroderma pigmentosum group D (XPD) helicases to unwind the DNA surrounding the damaged site. XPB will unwind the DNA in the 3′ to 5′ direction126 and XPD will unwind the DNA in the 5′ to 3′ direction127 from the site of the lesion. Once the DNA has been unwound, a patch of DNA ∼25–30 base pairs in length can be cleaved. The first incision is made on the 3′ side of the lesion by Xeroderma pigmentosum group G (XPG) endonuclease.128,129 This incision is made ∼3–5 base pairs to the 3′ side of the lesion130,131 and is required to be completed before a 5′ incision can be made.131 Excision repair cross-complementing 1 (ERCC1) and 4 (ERCC4) proteins will then form a complex and cleave the DNA phosphate backbone ∼22–25 base pairs to the 5′ side of the lesion.129,130,132 RPA remains bound throughout the incision step because it is required for facilitating release of the excised oligonucleotide.120 Following release of the oligonucleotide, proliferating cell nuclear antigen (PCNA) displaces the incision proteins,130 clearing the way for polymerase d or polymerase e to synthesize a new segment of DNA.131 The nicks in the phosphate backbone are then sealed by DNA ligase. Chromium-induced DNA crosslinks, as well as protein crosslinks involving a complete protein are thought to be the primary lesions obstructing DNA replication past guanine residues.132–134 Treatment of mammalian cells with nonlethal concentrations of Cr(VI) has been shown to greatly enhance recombination,135 lending support to the proposal that these types of crosslinks do form in cellular systems following Cr(VI) exposure. When a polymerase encounters one of these bulky chromiuminduced lesions, it will stall, causing the replication fork to collapse.136 Collapse of a replication fork creates a single-ended double-strand break.137 A variation of homologous recombination repair called replication fork repair is used to mend doublestrand breaks created by the collapse of a replication fork.138 As a side note, replication fork collapse can also be induced by the presence of an unrepaired single-strand break139 or by the presence of a rigid oxidized nucleobase lesion.96 Oxidation of the deoxyribose by high-valent chromium ions, as well as formation of secondary oxidative guanine lesions like Sp could thus trigger replication fork collapse if the subsequent single-strand breaks are not promptly repaired. Homologous recombination repair is also referred to as error-free repair,137 making it the most favourable form of recombination for repair of exogenous DNA damage. However, in the replication fork repair version of homologous recombination, sister chromatid exchanges can potentially arise if the leading strand template becomes covalently attached to the daughter lagging strand.139,140 Chromium compounds have been shown to induce sister chromatid exchanges,47–49 which may be partially attributed to homologous recombination repair following the collapse of replication forks at chromium-induced DNA and protein crosslinks.

17.6 Conclusions Even though chromium has been one of the most extensively researched metal carcinogens, there is still much that is not known about its fundamental mechanism of interaction with DNA. This current lack of understanding is due to the complexity

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Cr-Induced DNA Damage and Repair of ternary DNA adducts in Cr(III)-dependent mutagenesis; Biochem., 2001, 40, 549–560. Snow, E.T.; Xu, L.S.; Chromium(III) bound to DNA templates promotes increased polymerase processivity and decreased fidelity during replication in vitro; Biochem., 1991, 30, 11238–11245. Zhitkovich, A.; Importance of chromium-DNA adducts in mutagenicity and toxicity of chromium(VI); Chem. Res. Toxicol., 2005, 18, 3–11. Peterson-Roth, E.; Reynolds, M.; Quievryn, G.; Zhitkovich, A.; Mismatch repair proteins are activators of toxic responses to chromium-DNA damage; Mol. Cell. Biol., 2005, 25, 3596–3607. Reynolds, M.; Zhitkovich, A.; Cellular vitamin C increases chromate toxicity via a death program requiring mismatch repair but not p53; Carcinogenesis, 2007, 28, 1613–1620. Strauss-Soukup, J.K.; Vaghefi, M.M.; Hogrefe, R.I.; Maher III, L.J.; Effects of neutralization pattern and stereochemistry on DNA bending by methylphosphonate substitutions; Biochem., 1997, 36, 8692–8698. Jiricny, J.; Nyström-Lahti, M.; Mismatch repair defects in cancer; Curr. Opin. Genet. Dev., 2000, 10, 157–161. Modrich, P.; Lahue, R.; Mismatch repair in replication fidelity, genetic recombination, and cancer biology; Annu. Rev. Biochem., 1996, 65, 101–133 Kolodner, R.D.; Marsischky, G.T.; Eukaryotic DNA mismatch repair; Curr. Opin. Genet. Dev., 1999, 9, 89–96. Reynolds, M.; Stoddard, L.; Bespalov, I.; Zhitkovich, A.; Ascorbate acts as a highly potent inducer of chromate mutagenesis and clastogenesis: linkage to DNA breaks in G2 phase by mismatch repair; Nucleic Acids Res., 2007, 35, 465–476. Hirose, T.; Kondo, K.; Takahashi, Y.; Ishikura, H.; Fujino, H.; Tsuyuguchi, M.; Hashimoto, M.; Yokose, T.; Mukai, K.; Kodama, T.; Moden, Y.; Frequent microsatellite instability in lung cancer from chromate-exposed workers; Mol. Carcinog., 2002, 33, 172–180. Kunkel, T.; Erie, D.A.; DNA mismatch repair; Annu. Rev. Biochem., 2005, 74, 681–710. Modrich, P.; Mechanisms in eukaryotic mismatch repair; J. Biol. Chem., 2006, 281, 30305–30309. Salnikow, K.; Zhitkovich, A.; Genetic and epigenetic mechanisms in metal carcinogenesis and cocarcinogenesis: nickel, arsenic, and chromium; Chem. Res. Toxicol., 2008, 21, 28–44. Strauss, J.K.; Maher III, J.L.; DNA bending in asymmetric phosphate neutralization; Science, 1994, 266, 1829–1834. Crothers, D.M.; Upsetting the forces of balance in DNA; Science, 1994, 266, 1819–1820. Voitkun, V.; Zhitkovich, A.; Costa, M.; Complexing of amino acids to DNA by chromate in intact cells; Environ. Health Perspect., 1994, 102(3), 251–255. Zhitkovich, A.; Voitkun, V.; Costa, M.; Formation of amino acid-DNA complexes by hexavalent and trivalent chromium in vitro: importance of trivalent chromium and the phosphate group; Biochem., 1996, 35, 7575–7582. Nickoloff, J.A.; Heokstra, M.F.; DNA Damage and Repair, Vol. II: DNA Repair in Higher Eukaryotes, Humana Press Inc., Totowa, NJ, 1998, 350–355. Andrews, A.D.; Barrett, S.F.; Yoder, F.W.; Robbins, J.H.; Xeroderma pigmetosum neurological abnormalities correlate with colony-forming ability after ultraviolet radiation; Proc. Natl. Acad. Sci. USA, 1978, 75, 1984–1988. Burns, J.L.; Guzder, S.N.; Sung, P.; Prakash, S.; Prakash, L.; An affinity of human replication protein A for ultraviolet-damaged DNA; J. Biol. Chem., 1996, 271, 11607–11610. Clugston, C.K.; McLaughlin, K.; Kenny, M.K.; Brown, R.; Binding of human singlestranded DNA binding protein to DNA damaged by the anticancer drug cisdiamminedichloroplatinum (II); Cancer Res., 1992, 52, 6375–6379. Matsuda, T.; Saijo, M.; Kuraoka, I.; Kobayashi, T.; Nakatsu, Y.; Nagai, A.; Enjoji, T.; Masutani, C.; Sugasawa, K.; Hanaoka, F.; Yasui, A.; Tanaka, K.; DNA repair protein XPA binds replication protein A (RPA); J. Biol. Chem., 1992, 270, 4152–4157.

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123. Schaeffer, L.; Roy, R.; Humbert, S.; Moncollin, V.; Vermeulen, W.; Hoeijmakers, J.H.J.; Chambon, P.; Egly, J.M.; DNA repair helicase: a component of BTF2 (TFIIH) basic transcription factor; Science, 1993, 260, 58–63. 124. Sung, P.; Bailly, V.; Weber, C.; Thompson, L.H.; Prakash, L.; Prakash, S.; Human Xeroderma pigmentosum group D gene encodes a DNA helicase; Nature, 1993, 365, 852–855. 125. O’Donovan, A.; Scherly, D.; Clarkson, S.G.; Wood, R.G.; Isolation of active recombinant XPG protein, a human DNA repair endonuclease; J. Biol. Chem., 1994, 269, 15965–15968. 126. Matsunaga, T.; Mu, D.; Park, C.H.; Reardon, J.T.; Sancar, A.; Human DNA repair excision nuclease; J. Biol. Chem., 1995, 270, 20862–20869. 127. Huang, J.C.; Svoboda, D.L.; Reardon, J.T.; Sancar, A.; Human nucleotide excision nuclease removes thymine dimmers from DNA by incising the 22nd phosphodiester bond 5’ and the 6th phosphodiester bond 3’ to the photodimer; Proc. Natl. Acad. Sci. USA, 1992, 89, 3664–3668. 128. Mu, D.; Hsu, D.S.; Sancar, A.; Reaction mechanism of human DNA repair excision nuclease; J. Biol. Chem., 1996, 271, 8285–8294. 129. Park, C.H.; Bessho, T.; Matsunaga, T.; Sancar, A.; Purification and characterization of the XPF-ERCC1 complex of human DNA repair excision nuclease; J. Biol. Chem., 1995, 270, 22657–22660. 130. Nichols, A.F.; Sancar, A.; Purification of PCNA as a nucleotide excision repair protein; Nucleic Acids Res., 1992, 20, 2441–2446. 131. Hubscher, U.; Spadari, S.; DNA replication and chemotherapy; Physiol. Rev., 1994, 74, 259–304. 132. O’Brien, T.; Mandel, H.G.; Pritchard, D.E.; Patierno, S.R.; Critical role of chromium (Cr)-DNA interactions in the formation of Cr-induced polymerase arresting lesions; Biochem., 2002, 41, 12529–12537. 133. Bridgewater, L.C.; Manning, F.C.; Patierno, S.R.; Base-specific arrest of in vitro DNA replication by carcinogenic chromium: relationship to DNA interstrand crosslinking; Carcinogenesis, 1994, 15, 2421–2427. 134. Bridgewater, L.C.; Manning, F.C.; Patierno, S.R.; Arrest of replication by mammalian DNA polymerases a and b caused by chromium-DNA lesions; Mol. Carcinog., 1998, 23, 201–206. 135. Helleday, T.; Nilsson, R.; Jenssen, D.; Arsenic(III) and heavy metal ions induce intrachromosomal homologous recombination in the hprt gene of V79 Chinese hamster cells; Environ. Mol. Mutagen., 2000, 35, 114–122. 136. O’Brien, T.J.; Fornsaglio, J.L.; Ceryak, S.R.; Patierno, S.R.; Effects of hexavalent chromium on the survival and cell cycle distribution of repair-deficient S. cerevisiae; DNA Repair, 2002, 1, 617–627. 137. Helleday, T.; Lo, J.; van Gent, D.C.; Engelward, B.P.; DNA double-strand break repair: from mechanistic understanding to cancer treatment; DNA Repair, 2007, 6, 923–935. 138. Strumberg, D.; Pilon, A.A.; Smith, M.; Hickey, R.; Malkas, L.; Pommier, Y.; Conversion of topoisomerase I cleavage complexes on the leading strand of ribosomal DNA into 5’-phosphorylated DNA double-strand breaks by replication runoff; Mol. Cell. Biol., 2000, 20, 3977–3987. 139. Richardson, C.; Moynahan, M.E.; Jasin, M.; Double-strand break repair by interchromosomal recombination: suppression of chromosomal translocations; Genes Dev., 2000, 12, 3831–3842. 140. Richardson, C.; Jasin, M.; Frequent chromosomal translocations induced by DNA double-strand breaks; Nature, 2000, 405, 676–700.

18 Arsenic-Induced Carcinogenicity: New Insights in Molecular Mechanism Andrea Hartwig and Tanja Schwerdtle

18.1 Introduction Arsenic is a well-documented human carcinogen after both oral exposure and inhalation. While environmentally relevant arsenic species include inorganic as well as organic arsenicals, elevated cancer incidences have been attributed to inorganic arsenic, and research has focused on identifying the underlying mechanisms. A variety of inorganic arsenates or arsenites occur in water, soil and food, while the most common inorganic arsenical in the air is arsenic trioxide (As2O3). In general, trivalent arsenites tend to be more toxic than pentavalent arsenates; however, in many cases mixed exposure towards trivalent and pentavalent inorganic arsenic occurs and the precise chemical speciation is usually not known.1,2 Regarding worldwide public health, the most important medium for inorganic arsenic exposure is drinking water, causing many types of cancer and a wide range of further adverse health effects, including neurotoxicity, liver injury and the endemic ‘blackfoot disease’.3–6 Furthermore, occupational exposure is still relevant, even though the use of arsenicals has been restricted in industrialized countries considerably during recent years.

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

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18.2 Carcinogenicity and Cocarcinogenicity Predominant tumours in humans exposed chronically via inhalation or the gastrointestinal tract are lung and skin cancers, respectively. Additionally, case reports and epidemiological studies indicate increased incidences of tumours in the skin, the liver and the digestive tract after chronic arsenic inhalation, as well as increased risks of internal tumours (mainly bladder, lung, and to a lesser extent liver, kidney and prostate) after chronic arsenic ingestion (reviewed in references 3–6). Numerous scientific commissions and regulatory agencies have classified arsenic as a human carcinogen; among these are the International Agency for Research on Cancer (IARC), the German Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area (MAK commission) and the United States Environmental Protection Agency (USEPA).2,7–11 In contrast to humans, where the carcinogenic potential is clearly evident, experimental animals usually fail to demonstrate increased tumour incidences, with the exception of studies in mice demonstrating transplacental carcinogenesis.2,12–14 Nevertheless, cocarcinogenic effects have been observed in several experimental settings. Thus, early animal studies have shown that lung adenoma incidences were increased by benzo[a]pyrene and arsenic cotreatment, as compared to treatment with each chemical alone.15 More recently, arsenite has been demonstrated to enhance the carcinogenicity of UV radiation in mice, thus combined exposure to arsenite-enriched drinking water and solar UV radiation strongly increased skin tumour incidences, as compared to mice given UV alone.16,17 Similar cocarcinogenic effects were also seen in humans. The results from several epidemiological studies suggest more than additive effects on respiratory tract cancer incidences in smokers that were exposed to arsenic occupationally or via drinking water.18,19 Additionally, epidemiological studies provide some evidence that arsenite acts as a cocarcinogen in skin cancer (reviewed in reference 20). With respect to organic methylated arsenic metabolites, oral treatment of animals with dimethylarsinic acid (DMAV) has been shown to act as a tumour promoter in several organs and as a complete carcinogen in the rat bladder; however it is questionable whether the applied DMAV concentrations could be attained in vivo after exposure to nonacutely toxic inorganic arsenic doses (summarized in references 21 and 22).

18.3 Metabolism of Inorganic Arsenic After inhalation or ingestion of inorganic arsenic, humans and many other mammals have the ability to metabolize inorganic arsenic into organic arsenic forms that are more readily excreted via urine. After reduction of arsenate, arsenite is converted to its trivalent and pentavalent methylated metabolites, monomethylarsonous (MMAIII) and dimethylarsinous (DMAIII) acid, monomethylarsonic (MMAV) and dimethylarsinic (DMAV) acid. To date, the individual steps of biomethylation, which

Metabolism of Inorganic Arsenic Arsenite

Arsenate

OH O

AsV

OH

O-

HO

2e-

ATG

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O

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CH3 DMAV

GS

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CH3

CH3

CH3

CH3

CH3

DMAG

GSH

HO

AsIII DMAIII

CH3

O

AsV O-

CH3 DMAV

CH3

Figure 18.1 Biotransformation of inorganic arsenic into methylated metabolites. Shown are schemes of the originally proposed, generally accepted, pathway with successive oxidative methylation and reduction steps (A) and the recently proposed alternative pathway, involving formation of arsenic-glutathion complexes (B). AS3MT, arsenic (+3 oxidation state) methyltransferase; ATG, arsenite triglutathione; DMAG, dimethylarsinic glutathione; GSTw, glutathione S-transferase w; MADG, monomethylarsonic diglutathione; PNP, purine nucleoside phosphorylase; SAHC, S-adenosylhomocysteine; SAM, S-adenosylmethionine (summarized from references 2, 23 and 28)

catalyse conversion of inorganic arsenic to the methylated metabolites enzymatically, are still not fully understood (reviewed in references 23–26). Two possible metabolic pathways have been postulated, yielding the same methylated metabolites. First a pathway has been proposed with successive oxidative methylation and reduction steps.27 Recently, alternative metabolization steps have been discussed, focusing on glutathione (GSH) conjugation and subsequent methylation28 (Figure 18.1). Furthermore, recent work suggested that cysteine residues in cellular proteins might serve an analogous function in complexing the arsenic-containing substrate like GSH.29 Whereas biomethylation has long been thought to be a detoxification process, nowadays it is reasonable to conclude that some adverse health effects seen in humans chronically exposed to inorganic arsenic are in fact caused by these metabolites. With respect to the trivalent metabolites MMAIII and DMAIII, numerous recent

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studies have shown that they are as toxic, or even more toxic, compared to inorganic arsenic. Thus, with respect to molecular mechanisms, special emphasis is given to the biomethylated arsenicals, as they may at least contribute to inorganic arsenicinduced genotoxicity and presumably carcinogenicity (reviewed in references 30– 33). Whereas the ability of arsenic to induce carcinogenicity is beyond dispute, the mechanisms of how arsenic induces cancer in humans have not been clearly identified. Various modes of action have been proposed to be involved, which are discussed based on recent reviews22,26,30,32,34–37 and numerous original papers. Since evidence of inorganic arsenic-induced carcinogenicity in laboratory animals is mostly negative, modes of action studies are mostly conducted in cell culture systems.

18.4 Modes of Action 18.4.1 Induction of Genetic Damage Unlike many other carcinogens, arsenite neither directly interacts with DNA nor induces point mutations in bacterial or mammalian test systems (reviewed in references 4 and 10). However, arsenite has been shown to induce large deletion (multilocus) mutations in hamster human hybrid cells,38 micronuclei (MN) and chromosomal aberrations (CA), aneuploidy and sister-chromatid exchanges (SCE) in various mammalian cells (recently summarized in reference 2). Also in peripheral lymphocytes, oral mucosa or bladder urothelial cells of humans exposed towards elevated levels of arsenic in drinking water, numerous studies observed increases in chromosomal aberrations, micronuclei and sister chromatid exchanges (Table 18.1). In vitro studies in mammalian cells demonstrated the induction of DNA damage (strand breaks, oxidative base modifications, apurinic/apyrimidinic sites, DNA– protein crosslinks) and inhibition of DNA repair (see below). Thus, chromosomal alterations may be a secondary result of arsenite-induced DNA damage and DNA repair inhibition.32 Furthermore Li and Broome proposed a model, in which arsenite crosslinks tubulin and inhibits GTP binding, resulting in disturbed tubulin polymerization and mitosis,39 which may contribute to micronuclei formation. Additionally, arsenite and arsenate can cause gene amplification in mouse 3T6 cells.40 Since oncogenes are amplified in several animal and human tumours, the ability of arsenic to induce gene amplification may relate to its carcinogenic effects. Regarding the methylated species, MMAIII and DMAIII are generally genotoxic at lower concentrations than inorganic arsenic, while effects of the pentavalent metabolites MMAV and DMAV are either absent or restricted to much higher concentrations (e.g., references 41–45). In this context Kligermann and coworkers have recently summarized evidence that trivalent methylated arsenicals are potent clastogens and, due to spindle disruption, are aneugens as well.32 While arsenic is not a classical mutagen, it increases the genotoxicity, mutagenicity and clastogenicity of other DNA damaging agents, among others UV light,

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Table 18.1 Genotoxicity in humans after exposure towards elevated arsenic drinking water levels Individuals (exposed, controls) Country

Arsenic in drinking water

Effects in exposed individuals

Ref.

24 (11, 13) Mexico 437 (282, 155) Argentina 36 (18, 18) Nevada, USA 44 (22, 22) Argentina 69 (35, 34) Mexico 125 (70, 55) Chile 74 (42, 32) Finland 32 (19, 13) Inner Mongolia, China 66 (45, 21) West Bengal, India 95 (59, 36) West Bengal, India 317 (163, 154) West Bengal, India 320 (165, 155) West Bengal, India 217 (106, 111) North Chile 206 (105, 102) North Chile 306 (204, 102) West Bengal, India

controls: 26 µg/l exposed: 390 µg/l controls: 20 µg/l exposed: 130 µg/l controls: 16 µg/l exposed: 1313 µg/l controls: 8.4 µg/l exposed: 205 µg/l controls: 30 µg/l exposed: 408 µg/l controls: 15 µg/l exposed: 600 µg/l controls: 7 µg/l exposed: 410 µg/l controls: 4.5 µg/l exposed: 527.5 µg/l controls: 5.5 µg/l exposed: 368.1 µg/l controls: 6.4 µg/l exposed: 211.7 µg/l controls: 9.2 µg/l exposed: 214.7 µg/l controls: 9.1 µg/l exposed: 215 µg/l controls: 2.0 µg/l exposed: 750 µg/l controls: 2.0 µg/l exposed: 750 µg/l controls: 7.2 µg/l exposed: 220 µg/l

L: CA (↑), SCE (-), mutations in HPRT-locus ↑ L: SCE ↑

53 54

UC: MN (↑)

55

L: MN ↑, trisomy, SCE (-) L: CA (↑), OC,BC: MN (↑) UC: MN (↑)

56

58

L: CA ↑

59

OC,UC: MN ↑

60

L, OC, UC: MN ↑

61

L: SCE ↑, CA ↑

62

L,OC,UC: MN ↑

63

L: CA ↑, SCE ↑

64

L: MN ↑

65

OC: MN (↑)

66

L,OC,UC: MN ↑ L: CA ↑

67

57

CA, chromosomal aberrations; L, lymphocytes; MN, micronuclei; OC, oral mucosa cells; SCE, sister chromatid exchanges; UC, urothelial cells; ↑, significant increase; (↑), increase but not significant, (-), no effect

benzo[a]pyrene and alkylating agents,46–51 which may be explained by the interference with DNA repair processes.52 This is consistent with the comutagenic effect of arsenic, resulting in arsenic cocarcinogenesis, which has been shown in vivo (see below). 18.4.2 Involvement of Reactive Oxygen (ROS) and Nitrogen Species (RNS) in Arsenic Response Numerous studies provide strong evidence that oxidative stress mediated by increased levels of ROS and RNS is an important molecular mechanism contributing to arsenic-induced genotoxicity and carcinogenicity (reviewed in references

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35, 68 and 69). Thus, in diverse cellular systems, arsenite increases the generation of superoxide anions ( iO−2 ) and hydrogen peroxide (H2O2), and modulates the level of nitric oxide (NO). Subsequently these reactive species can be converted to other, more damaging reactive species, such as the hydroxyl radical (·OH) and peroxynitrite (ONOO−). Several origins of elevated levels of reactive species have been suggested. They include interactions with the respiratory chain,70,71 their generation during metabolism of inorganic arsenic, such as the formation of intermediary dimethylarsine and radical arsenic species,72–79 the release of iron from ferritin by dimethylated arsenic species,80,81 and modulation of NO synthases.31,68,82 All of these mechanisms may lead to oxidative stress, resulting from an imbalance between free radical generation and cellular antioxidant defence systems.83 Moreover, arsenic has not only been shown to increase the generation of reactive species, but also to interact with cellular protection mechanisms. Thus arsenic is believed to change cellular redox homeostasis by decreasing cellular GSH. This might be due to the ability of trivalent arsenicals to complex with thiol groups, resulting in GSH binding and depletion, consumption of GSH during arsenic metabolism, as well as effects of trivalent methylated arsenicals on glutathione-related enzymes.84,85 With respect to genotoxicity, the application of radical scavengers revealed the involvement of arsenite-induced ROS and RNS in the induction of lipid peroxidation, protein oxidation, DNA damage (summarized in reference 35) and DNA repair inhibition.86 Furthermore chronic low-dose arsenic alters gene expression and protein levels that are associated with oxidative stress and inflammation (e.g., references 87–89), which may in part be due to oxidation of major transcriptional redox sensitive regulators (e.g., Nrf2, nuclear factor-erythroid 2-related factor 2) of altered genes. Thus the induced reactive species are known to activate signal cascades such as the mitogen-activated protein kinases (MAPKs) cascade and the transcription factors AP-1 (activator protein-1) and NFkB (nuclear factor-kB) (summarized in references 31 and 90). Finally, the fact that arsenite-induced oxidative stress is also evident in populations exposed to arsenic-contaminated drinking water underscores its role in arsenic-induced carcinogenicity (see, for example, references 91–97).

18.4.3 Epigenetic Effects In addition to the induction of genetic damage, altered DNA methylation may also contribute to carcinogenicity. Thus, in tumours global methylation is typically reduced, but some gene-specific promoter methylation is increased, thereby affecting the expression of protooncogenes and tumour suppressor genes, resulting in cell transformation and upregulated cell growth.98,99 Over the last 10 years accumulating evidence from cell culture studies, experimental animals and also from arsenicexposed humans suggests that epigenetic changes contribute to arsenic-induced carcinogenicity. Thus both hypo- and hypermethylation have been observed after arsenic exposure. For instance, increased cytosine methylation in the p53 promoter was detected in A549 human lung cells by arsenite and arsenate, but not by DMAv 100

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(further effects on p53 are discussed below). After exposure of A549 human lung and UOK human kidney cells to arsenite for several weeks, Zhong and Mass observed both hypo- and hypermethylation of different genes,101 whereas in a rat liver cell line102 and human HaCaT keratinocytes103 global DNA hypomethylation was found following chronic exposure to low levels of arsenite. In A/J mice, which were chronically orally exposed to arsenate and developed lung tumours, hypermethylation was observed with the consequence of diminished p16INK4a and RASSF1A expression occurring in these tumours.104 With respect to humans, dosedependent hypermethylation of the p53 and p16 genes was observed in blood samples of arsenic-exposed skin cancer patients in West Bengal.105 Additionally, in a population-based study of human bladder cancer, arsenic exposure measured as toenail arsenic was associated with promoter hypermethylation of the tumour suppressor proteins RASSF1A and PRSS3, but not p16INK4a.106 While hypomethylation may be due to inhibition of DNA methyltransferases or depletion of the methyl donor SAM,102,103 which is a common cofactor in DNA and arsenic methylation, hypermethylation is not easily understood and further studies are required to resolve this question.

18.4.4 Inhibition of DNA Repair To ensure faithful duplication and inheritance of genetic material, organisms have evolved to efficiently detect and remove DNA damage resulting from either endogenous sources (cellular metabolic processes) or exogenous sources (environmental factors). Dysregulation of DNA damage-response processes, including cell cycle arrest, DNA repair, senescence or apoptosis, contribute to genomic instability on the cellular level, predisposing the organism to cancer.107 Therefore, it is absolutely essential for cells to efficiently respond to DNA damage through coordinated and integrated cell cycle checkpoints and DNA repair pathways.108–110 Concerning DNA repair inhibition as one underlying mechanism of arsenic-induced carcinogenicity, several studies point to an interaction of arsenic with various DNA repair pathways, including nucleotide (NER) and base excision repair (BER) (reviewed in references 30 and 34). NER is capable of removing a wide variety of bulky, DNA-helix-distorting lesions, caused by UV radiation, chemotherapeutic drugs or environmental mutagens. During the last few years there has been accumulating evidence that low, noncytotoxic concentrations of arsenite inhibit nucleotide excision repair (NER),47,86,111–115 which is even more pronounced with the trivalent methylated metabolites MMAIII and DMAIII.42,116 Thus, arsenic has been shown to decrease removal of bulky DNA adducts induced by UV radiation47,86,111–113,115 or benzo[a]pyrene in cultured cells and laboratory animals.42,114,116,117 NER works through a ‘cut-andpatch’ mechanism; after DNA damage recognition, incisions at sites flanking the lesion occur and an oligonucleotide (24–32 nucleotides) containing the lesion is excised, followed by subsequent restoration of the original DNA sequence by polymerisation/ligation using the nondamaged strand as a template. In total, about

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30 proteins are involved (recently summarized in references 108 and 118). In the first stage, arsenite seems to disturb the DNA damage recognition/incision step, but effects on DNA ligation are observed as well113 One possible mechanism of arsenic-induced toxicity may lie in its ability to react with thiols, for example in zinc-binding structures prevalent in many transcription factors, cell cycle control and DNA repair proteins (recently reviewed in reference 119). In case of the central NER DNA damage recognition protein XPA, trivalent arsenicals have been shown to release zinc from a 37 amino acid XPAzf peptide, representing the zinc-finger domain of the human XPA protein.42 Thus equimolar MMAIII released ZnII easily, forming mono- and diarsenical derivatives of XPAzf.120 In contrast, a 10-fold excess of arsenite was required to partially oxidize the zinc-finger structure of XPAzf. Thus, zinc-binding structures may be sensitive targets for arsenicals, even though the actual species involved in the specific interaction differ. With respect to XPA function, subcellular studies observed no or only a slight decrease on XPA binding to UVC-121 or MMC-damaged122 oligonucleotides by arsenite. To date the most sensitive target related to DNA repair is inhibition of poly(ADP-ribosyl)ation by trivalent arsenicals at nanomolar concentrations, with MMAIII and DMAIII showing the strongest effects.123,124 Poly(ADP-ribosyl)ation, which is mediated mainly by the poly(ADP-ribose) polymerase 1 (PARP-1), plays a major role in DNA damage signalling and the recruitment of DNA repair proteins to sites of damage. Although the role of PARP-1 is not fully understood as yet, there is strong evidence that PARP-1 contributes to base excision repair (BER),125 which among others is responsible for the repair of oxidative DNA damage, and perhaps also to NER. In this context, Poonepalli et al. reported that in mouse embryonic fibroblasts, a lack of PARP-1 gene product enhances cellular sensitivity to arsenite.126 With respect to BER, arsenite has also been shown to inhibit DNA ligase III activity in cells and nuclear extracts, most probably indirectly by altering cellular redox levels or by affecting signal transduction pathways related to ligase activity.127,128 As mentioned before, effects on the tumour suppressor protein p53 may also contribute to arsenic-induced carcinogenicity. Following activation by posttranslational modifications such as phosphorylation, acetylation and poly(ADP-ribosyl)ation, p53 plays a guarding role in maintaining genome integrity by activating the transcription of genes involved in cell cycle check points, cellular senescence, apoptosis and DNA repair (e.g., XPC, p48, hOGG1). At present effects of arsenic on p53 are not well understood, since responses are different depending on cell type and incubation times. Investigations in human fibroblasts and human lymphoblastoid cell lines have identified that arsenite could induce p53 accumulation through an ATMdependent pathway.129,130 On the other hand, arsenite and MMAIII have been shown to inhibit p53 phosphorylation and to reduce overall p53 level, as well as p53 DNAbinding activity.116,131 Furthermore a recent study has postulated increased formation of conformationally mutated p53 by arsenite.132 Due to the loss of its zinc-containing folded wild-type conformation, this conformationally mutated p53 is no longer capable of function as a transcription factor;133,134 many cancer-associated mutations cause loss of this conformation.135

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Besides direct and indirect interaction with repair or repair-associated proteins, arsenic compounds may also diminish DNA repair by altering the expression of specific DNA repair genes. Thus Hamadeh et al.136 and Andrew et al.137 reported a decreased gene expression of various repair genes in arsenite-exposed human normal keratinocytes and cultured human bronchial epithelial cells, respectively. Our recent work demonstrates that arsenite strongly decreased expression and protein level of XPC, which is believed to be the principle initiator of global genome NER. This phenomenon was even more pronounced with MMAIII. This led to diminished association of XPC to sites of local UVC damage in human fibroblasts, resulting in decreased recruitment of further NER proteins and therefore disrupted DNA repair.138 In humans, arsenic exposure via drinking water was correlated in a dosedependent manner to decreased expression of NER and BER genes and diminished repair of lesions in lymphocytes.139,140 Additionally, a recent study showed that upon induction of DNA damage by gamma irradiation, the repair capacity in whole blood of arsenic-exposed individuals with premalignant hyperkeratosis was significantly less, compared to that of individuals without skin lesions.141 Table 18.2 summarizes studies providing evidence for inhibition of nucleotide and base excision repair by inorganic and methylated arsenicals.

18.5 Conclusions and Research Needs Taken together, the carcinogenicity of inorganic arsenic is well documented, but underlying mechanisms are not completely understood. Current evidence suggests that indirect mechanisms play a predominant role: rather than direct binding to DNA, interactions with proteins are of major importance, leading to oxidative stress and modulating the cellular response to DNA damage, which may in turn decrease genomic integrity (Figure 18.2). This would also explain the enhancement of benzo[a]pyrene and UV-induced carcinogenicity in animals and exposed humans, which may be due to the disturbance of DNA repair processes. However, in spite of many advances in arsenic-related toxicology, several aspects need to be addressed in future research. They include further insights into arsenic metabolism in humans and the relevance of metabolites in arsenic-induced genotoxicity and carcinogenicity. Furthermore, the impact of genetic polymorphisms, both in arsenic biomethylation and in DNA repair genes, needs to be clarified. Finally, the toxicological impact of organic arsenic compounds such as arsenosugars and arsenolipids warrants special attention.

Acknowledgements This work was supported by the DFG grant number Ha 2372/3–4 and SCHW 903/3–2.

Arsenite (100 µM) MMAIII (10 µM) DMAIII (10 µM) MMAV (5 mM) DMAV (no effect up to 10 mM)

Arsenite (5 µM) MMAIII (2.5 µM) DMAIII (2.5 µM) MMAV (250 µM) DMAV (250 µM) Arsenite (5 µM) MMAIII (1 µM) DMAIII (1 µM) Arsenite (500 µM)

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Arsenic compound (lowest effective dose)

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h. CRL2522 fibroblasts h. colon HCT116 cancer cells

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h. lymphocytes SV40-transformed fibroblasts (MRC5CV1) h. fibroblast CHO cells h. fibroblast Sprague-Dawley rats h. TK6 lymphoblastoid cells

CHO-K1 cells

h. fibroblast (SF24, XP6KO, XP2SA) V79 cells

Species (test system)

removal of BPDE-induced DNA adducts ↓ impairment of p53 induction in response to BPDE treatment binding of XPA to the MMC C-DNA interstrand crosslink ↓ release of zinc from XPAzf

NER NER

Removal of B[a]P-induced DNA adducts ↓ removal of UV-induced photolesions ↓ comutagenicity with UV removal of BPDE-induced DNA adducts ↓

NER

NER

NER

NER

NER NER

NER

NER

NER

Pathway

Removal of UV-induced photolesions ↓ Removal of UV-induced photolesions ↓

removal of UV-induced photolesions ↓ coclastogenicity with UV Removal of B[a]P-induced lesions ↓

comutagenicity with UV

removal of UV-induced photolesions ↓

End Point

Table 18.2 Studies providing evidence for inhibition of nucleotide and base excision repair by arsenicals

42, 120

122

116

42

114 115

113 86

117

111, 112

47

Ref.

500 Arsenic-Induced Carcinogenicity

h. BEAS-2B bronchial epithelial cells h. lymphocytes (in vivo) h. lymphocytes (in vivo) (in vitro) h. VH10hTert fibroblasts

Arsenite (5 µM)

H2O2-stimulated poly(ADP-ribosyl)ation ↓

PARP-1 activity ↓

h. Molt-3 T-cell lymphoma cells

h. HeLa S3 cervic carcinoma cells

h. HeLa S3 cervic carcinoma cells

PARP-1

Arsenite (0.01 µM)

MMAIII (0.001 µM) DMAIII (0.001 µM) MMAV/DMAV (no effect up to 250 µM) Arsenite (10 µM) MMAIII (10 µM) DMAIII (10 µM)

BER (NER)

BER (NER) BER (NER) BER (NER)

BER

BER BER

NER

NER BER NER BER NER NER

Pathway

124

124

123

142

89

127 128

138

140 139

137

136

Ref.

2-AAAF, 2-acetoxyacetylaminofluorene; APE1, AP-endonuclease 1, BER endonuclease; B[a]P, benzo[a]pyrene; BER, base excision repair; BPDE, benzo[a]pyrene diolepoxide; ERCC1, excision repair cross-complementing rodent repair deficiency complementation group 1; FEN1, FLAP endonuclease 1; h, human; LIG1, DNA-Ligase 1; MMC, mitomycin C; NER, nucleotide excision repair; PARP-1, poly(ADP-ribose)polymerase-1; PCNA, proliferating cell nuclear antigen; p48, subunit of the DDB2 NER damage recognition protein; Pold/e, DNA polymerase d/e; RFC, replication factor C; UDG2, uracil DNA glycosylase 2; XP, Xeroderma Pigmentosum group proteins, NER damage recognition / incision proteins; XPAzf, peptide representing the zinc finger domain of the human XPA protein.

H2O2-stimulated poly(ADP-ribosyl)ation ↓

BER incision activity (cell extracts) ↓ APE1↓, Polb↓, Polb↓, PARP-activity ↓

Arsenite (2.5 µM)

Arsenite (≥5 µM)

Chinese V79 hamster cells h. HOS osteosarcoma cells h. AG06 keratinocytes LIG1 h. HaCaT keratinocytes

XPC↓, p48↓, p53↓ FEN1↓, Pold↓, Pole↓, UDG2↓, p53↓ XPC ↓, XPD ↓, RFC ↓, PCNA ↓, LIG1↓ APE1↓, LIG1↓ ERCC1↓, XPF ↓, XPB↓ ERCC1↓, ERCC1 protein ↓ repair of 2-AAAF-induced lesions ↓ Disturbed assembly of NER proteins after UV-irradiation XPC ↓, p48↓, XPC protein ↓ Nuclear DNA ligase activity ↓ Ligase activity in nuclear extracts of cells ↓

End Point

Arsenite (10 µM) Arsenite (10 µM) (>2 mM)

Arsenite (5 µM) MMAIII (0.1 µM)

As in drinking water As in drinking water

h. normal epidermal keratinocytes

Species (test system)

Arsenite (0.5 µM)

Arsenic compound (lowest effective dose)

Acknowledgements 501

502

Arsenic-Induced Carcinogenicity Exposure towards arsenicals Metabolism in vivo

Inorganic and organic trivalent and pentavalent arsenicals

Oxidative stress, protein binding, altered DNA methylation

• DNA damage, protein oxidation, lipid peroxidation

• Alteration of gene expression

• Inhibition of DNA repair

• Modulation of signal transduction pathways

Persistence of DNA lesions, mutations, chromosomal damage, upregulation of stress proteins and protooncogenes, downregulation of tumour suppressor genes, modulation of cell cycle

Accumulation of mutations, enhanced cell poliferation

Carcinogenicity, cocarcinogenicity

Figure 18.2 Major mechanisms in arsenic induced carcinogenicity

Abbreviations 2-AAAF AP-1 APE1 AS3MT ATG B[a]P BER BPDE CA DMAV DMAIII DMAG GSH GSTw ERCC1

2-acetoxyacetylaminofluorene activator protein-1 AP-endonuclease 1 arsenic (+3 oxidation state) methyltransferase arsenite triglutathione benzo[a]pyrene base excision repair benzo[a]pyrene diolepoxide chromosomal aberrations dimethylarsinic acid dimethylarsinous acid dimethylarsinic glutathione glutathione glutathione S-transferase w excision repair cross-complementing rodent repair deficiency complementation group 1

References

FEN1 h IARC L LIG1 MADG MAK commission MMAV MMAIII MN MMC NER NFkB Nrf2 OC PARP-1 PCNA p48 PNP Pold/e RFC RNS ROS SAHC SAM SCE UC UDG2 USEPA XP XPAzf

503

FLAP endonuclease 1 human International Agency for Research on Cancer lymphocytes DNA ligase 1 monomethylarsonic diglutathione German Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area monomethylarsonic acid monomethylarsonous acid micronuclei mitomycin C nucleotide excision repair nuclear factor-kB factor-erythroid 2-related factor 2 oral mucosa cells poly(ADP-ribose)polymerase-1 proliferating cell nuclear antigen subunit of the DDB2 NER damage recognition protein purine nucleoside phosphorylase DNA polymerase d/e replication factor C reactive nitrogen species reactive oxygen species S-adenosylhomocysteine S-adenosylmethionine sister-chromatid exchanges urothelial cells uracil DNA glycosylase 2 United States Environmental Protection Agency Xeroderma pigmentosum group proteins, NER damage recognition/incision proteins peptide representing the zinc finger domain of the human XPA protein

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128. Hu, Y.; Su, L.; Snow, E.T.; Arsenic toxicity is enzyme specific and its affects on ligation are not caused by the direct inhibition of DNA repair enzymes; Mutat. Res. 1998, 408, 203–218. 129. Yih, L.H.; Lee, T.C.; Arsenite induces p53 accumulation through an ATM-dependent pathway in human fibroblasts; Cancer Res. 2000, 60, 6346–6352. 130. Menendez, D.; Mora, G.; Salazar, A.M.; Ostrosky-Wegman, P.; ATM status confers sensitivity to arsenic cytotoxic effects; Mutagenesis 2001, 16, 443–448. 131. Tang, F.; Liu, G.; He, Z.; Ma, W.Y.; Bode, A.M.; Dong, Z.; Arsenite inhibits p53 phosphorylation, DNA binding activity, and p53 target gene p21 expression in mouse epidermal JB6 cells; Mol. Carcinog. 2006, 45, 861–870. 132. Chai, C.Y.; Huang, Y.C.; Hung, W.C.; Kang, W.Y.; Chen, W.T.; Arsenic salt-induced DNA damage and expression of mutant p53 and COX-2 proteins in SV-40 immortalized human uroepithelial cells. Mutagenesis 2007, 22, 403–408. 133. Butler, J.S.; Loh, S.N.; Structure, function, and aggregation of the zinc-free form of the p53 DNA binding domain; Biochemistry 2003, 42, 2396–2403. 134. Wang, P.L.; Sait, F.; Winter, G.; The ‘wildtype’ conformation of p53: epitope mapping using hybrid proteins; Oncogene. 2001, 20, 2318–2324. 135. Martin, A.C.; Facchiano, A.M.; Cuff, A.L.; Hernandez-Boussard, T.; Olivier, M.; Hainaut, P.; et al., Integrating mutation data and structural analysis of the TP53 tumor-suppressor protein; Hum. Mutat. 2002, 19, 149–164. 136. Hamadeh, H.K.; Trouba, K.J.; Amin, R.P.; Afshari, C.A.; Germolec, D.; Coordination of altered DNA repair and damage pathways in arsenite-exposed keratinocytes; Toxicol. Sci. 2002, 69, 306–316. 137. Andrew, A.S.; Warren, A.J.; Barchowsky, A.; Temple, K.A.; Klei, L.; Soucy, N.V.; et al., Genomic and proteomic profiling of responses to toxic metals in human lung cells; Environ. Health Perspect. 2003, 111, 825–835. 138. Nollen, M.; Ebert, F.; Moser, J.; Mullenders, L.H.; Hartwig, A.; Schwerdtle, T.; Impact of arsenic on nucleotide excision repair; XPC function, protein level and gene expression; Molecular Nutrition and Food Research, 2008, submitted. 139. Andrew, A.S.; Burgess, J.L.; Meza, M.M.; Demidenko, E.; Waugh, M.G.; Hamilton, J.W.; et al., Arsenic exposure is associated with decreased DNA repair in vitro and in individuals exposed to drinking water arsenic; Environ. Health Perspect., 2006, 114, 1193–1198. 140. Andrew, A.S.; Karagas, M.R.; Hamilton, J.W.; Decreased DNA repair gene expression among individuals exposed to arsenic in United States drinking water; Int. J. Cancer, 2003, 104, 263–268. 141. Banerjee, M.; Sarma, N.; Biswas, R.; Roy, J.; Mukherjee, A.; Giri, A.K.; DNA repair deficiency leads to susceptibility to develop arsenic-induced premalignant skin lesions; Int. J. Cancer, 2008, 123, 283–287. 142. Yager, J.W.; Wiencke, J.K.; Inhibition of poly(ADP-ribose) polymerase by arsenite; Mutat. Res., 1997, 386, 345–351.

Index

Page numbers in italics refer to figures and diagrams. adenine dirhodium complexes, interaction with, 303, 304–305 metal ion affinities, 3 metal-DNA supramolecular chemistry 1D coordination polymers, 110– 113, 112, 115, 115–116, 116, 117 2D coordination polymers, 118– 119, 119 3D coordination polymers, 119– 121, 120, 121 chelates, 124, 125 H-bonding patterns, 123, 124 triangular supramolecules, 100–101 organotin complexes, interaction with, 309 platinum complexes, interaction with, 138, 138–139, 217, 220–222, 222 A-DNA, 335 affinity chromatography, 337–338 ammonium DNA affinity element, use as, 383 G-quadruplexes competition with Na+ and K+, 75–76

coordination in d[(G4T4G4)]2, 74 movement within, 79–80 residence times, 78–79 role in, 59 structure, effect on, 77–78 aptamers, 55, 275–276, 398 ARCUT system, 385, 385 arsenic carcinogenicity and cocarcinogenicity, 492, 502 exposure pathways, 491 genotoxicity, 495 metabolism of inorganic arsenic, 492–494, 493 modes of cancer induction, proposed, 502 epigenetic effects, 496–497 induction of genetic damage, 494–495 inhibition of DNA repair, 497–499, 500–501 oxidative stress, 495–496 ascorbate, 465–467, 466, 479 barium, 59, 60 base excision repair, 178, 179, 180, 193

Metal Complex–DNA Interactions Edited by Nick Hadjiliadis and Einar Sletten © 2009 Blackwell Publishing Ltd. ISBN: 978-1-405-17629-3

512

Index

effected by arsenic, 498 repair of Cr(VI) induced nucleobase lesions, 473, 473–474 B-DNA, 22–23, 238, 239, 335 biomethylation of inorganic arsenic, 492–494, 493 cadmium metal-DNA supramolecular chemistry 1D coordination polymers, 113, 114, 114–116, 115, 116 supramolecular squares, 102, 102– 103, 104 caesium, 59 calcium basic properties of Ca2+ cations, 416–417 Ca2+ in substrate binding, 422–423 G-quadruplexes coordination in d(TG4T), 71, 72 role in, 59–60 polymerases and nucleases, inhibition of, 425 preferred coordination geometry of Ca2+ cations, 417 cancer, modes of action, 236 arsenic-induced cancers, 494–499, 502 base mismatches, 307 chromium-induced cancers, 464–465 telomere dysfunction, 209, 225 cancer therapies, 236. see also photodynamic therapy (PDT) antitelomerase drugs, 213, 223 chemotherapy, 176 oligonucleotide therapies, 273–276 platinum drugs, 135–137, 136, 200, 216–217, 224–225 potential G-quadruplex targets for antitumour drugs, 215–216 ruthenium complexes, 339 telomerase inhibition, 213, 225 carboplatin, 135, 136, 188 catalysis in nucleic acid enzymes. see nucleic acid enzyme catalysis cerium Ce2-HXTA complex in DNA hydrolysis, 378 Ce(IV) complexes in DNA hydrolysis, 373–374, 383–384

Ce(IV)-EDTA complex in ARCUT system, 385, 385 chemical shifts C chemical shifts, 446 calculation, 452–453 N chemical shifts, 447, 450, 452 in T-HgII-T base pair formation, 452–454 chromium adduct formation, 477–478, 478 carcinogenic classification and limits, 463–464 carcinogenic properties, 464–465, 479, 479–480 chromate structure, 464 chromium(III) structure, 464 clastogenic classification, 476 Cr(VI) reductions, intracellular by ascorbate, 465–467, 466 by glutathione, 466, 467 by hydrogen peroxide, 466, 467 Cr(VI)-induced DNA crosslinks formation, 480–481, 481 repair, 481–482 DNA damage, 468–473, 480–482 repair, 473–474, 476–477, 478–480, 479, 481–482 exposure sources, 468 hydrogen chromate structure, 464 industrial uses, 464 lesions formed, 472 8-oxoG, 469, 469–470, 470 crosslinks, 480–482 guanidinohydantoin lesions, 471, 472 spiroiminodihydantoin lesions, 470–471, 471, 472 nucleobase oxidation electron abstraction pathway, 469–473 reactive oxygen species pathway, 468–469 reduction potentials, 469 repair, 473–474 sugar oxidation C1′ hydrogen abstraction mechanism, 475, 475 C4′ hydrogen abstraction mechanism, 475–476, 476 repair, 476–477

Index cisplatin (and related platinum complexes), 135, 136. see also platinum drugs binding patterns, 18 conformer distribution in cisplatin adducts of G derivatives adducts with 1-Me-5′-GMP, 154–155 adducts with 3′-GMP, 152–154, 153 adducts with 5′-GMP, 154 conformer distribution signature, 152, 153 quantification of atropisomer distribution, 156 cytotoxic activity, 301–302 DNA adducts repair, 193–200 repair, inhibition of, 183 structure, 188–189 types, 139, 139–140 DNA adducts with tethered guanine bases chirality, 158 flexibility, 161–163, 162 interligand interactions, 159 multiple conformers, 159–160 structure of d(GpG), 157 wrapping of platinum binding site, 160–161, 161 G-quadruplex structures, interaction with human telomeric sequence AG3(T2AG3)3, 220, 220–221 known classes, 217–218, 218 platinum-acridine complex, 221– 222, 222 platinum-quinacridine complex, 221–222, 222 Tetrahymena sequence (T2G4)4, 218–219, 219 method of anticancer activity, 216–217, 301 models of G/G crosslinks, 140–141 platinated oligonucleotides, 279, 280, 287 retro modelling, 141–145, 142, 144 structure, 136 telomerase, interaction with, 223–224 telomeric DNA, interaction with, 223, 224

513

cobalt Co(II)-dependent DNAzymes, selection of, 398–399, 399 Co(III) complexes in DNA hydrolysis, 374, 383 G-quadruplexes, role in, 59–60 oligodeoxynucleotides, interaction with, 7, 15 copper copper complex-peptide conjugate, 353–354 Cu-(GMP) complex, 4, 5 Cu(II) complexes in DNA hydrolysis, 375–377, 376 DNA melting temperature, effect on, 4, 16 DNAzyme for Cu(II), 396 DNAzyme-based Cu2+ sensor, 406 metal-DNA supramolecular chemistry, 107 1D coordination polymers, 110– 113, 112 2D coordination polymers, 118, 118 3D porous coordination polymer, 119–120, 120 oligodeoxynucleotides, adducts with, 15–16 crosslinking of oligonucleotides, 276–278, 277, 278, 289–295 cytosine depurination, 46–47 double-proton transfer, 44–46, 45 metal ion affinities, 3 metal-DNA supramolecular chemistry 1D coordination polymers, 110, 110, 113 2D coordination polymers, 118, 118 chelates, 124 H-bonding patterns to metals, 122 large supramolecules, 107–108, 108 square supramolecules, 104 triangular supramolecules, 101, 101–102 platinum binding sites, possible, 138, 138, 140 point defects, 48 depurination, 46–47, 48 dirhodium complexes, 303–305, 304

514

Index

DNA. see also B-DNA; M-DNA catalytic properties, 31 control of metal complexes, 124, 125 DNA binding. see also metal ion-DNA interactions, binding chromium, 477–478, 478 groove binding, 22–26 intercalation vs. groove binding, 359–361 mismatch recognition, 308 nucleobase-metal binding angles, 97 photobinding, 259–260 platinum, 138, 138–140, 139 ruthenium, 325–329. see also ruthenium, dinuclear complexes as DNA probes; ruthenium, mononuclear complexes as DNA probes stability constants, 34–38, 35, 36, 37, 38 transition metal polyazine complexes, 257–258 DNA bulge sequences, 330, 330–331, 331, 333 DNA cleavage degradation pathways, 370, 370–371 hydrolysis. see DNA hydrolysis oxidative vs. hydrolytic, 362 phosphoryl transfer reaction, 415–416, 416 photocleavage, 6, 7–9, 260–262, 305, 308, 352 DNA damage. see also DNA photomodification and photochemical degradation agents of damage, 175, 176 arsenic-induced, 494–502 chromium-induced, 468–473, 480–482 cisplatin-induced, 140 deletion, 48 double-strand breaks, 182, 199 mismatches, 307–308 oxidation, 308 persistence, factors contributing to, 185 photosensitizer damage, 255 platinum adducts, 176–177 types of damage, 175–176 DNA grooves intercalation vs. groove binding, 359–361 sequence specific binding divalent cations, 24–26 general binding trends, 26 monovalent cations, 23–24 structure, 22–23

DNA hairpin sequences, 332–334, 333 DNA hydrolysis ARCUT system, 385, 385 metallic hydrolytic agents bimetallic complexes, 377–380 conjugated with DNA affinity subunits, 380–384 conjugated with sequence-selective elements, 384–385 free ions, 373–374 metal ions, 371, 386 mononuclear complexes, 374–377 reaction pathway, 370, 370–371 research aims, 369–370 research considerations bimetallic systems, development of, 388 DNA affinity, increasing, 388–389 hydrolytic efficiency, 386–388, 387 ideal features of hydrolytic agents, 386 DNA photomodification and photochemical degradation mechanism of photomodification, 257–258 mechanisms of photochemical degradation photobinding DNA through a coordinated ligand, 260 photobinding DNA through a metal centre, 259 photocleavage of DNA, 260–262, 262 DNA polymerases, 179, 418, 418 DNA recognition code, 348, 348–349 DNA repair arsenic-induced inhibition, 494, 497–499 defined, 176 double-stranded breaks, repair of, 182–184, 184, 184–185, 186 implications for design of platinum drugs, 200 repair proteins and antitumour platinum compounds, 189 base excision repair system, 189 high-mobility group proteins, 193 homologous recombination system, 192 mismatch repair system, 191–192 nonhomologous end-joining system, 192

Index nucleotide excision repair system, 189–191 translesion synthesis DNA polymerases, 192–193 single-strand breaks, repair of, 476–477 types base excision repair, 178, 179, 180, 193, 473, 473–474 direct reversal of damage, 177–178 DNA damage bypass (translesion synthesis), 185–188, 187 homologous recombination, 184– 185, 186, 199–200 mismatch repair, 180–182, 183, 198–199, 478–480 nonhomologous end-joining, 199 nucleotide excision repair, 178–180, 181, 193–198, 481–482 recombination repair, 482 translesion synthesis (DNA damage bypass), 185–188, 187 DNA synthesis, 415, 416 DNA-metal interactions. see metal complexDNA interactions; metal ionDNA interactions DNAzymes catalytic activities, 395 cofactors, 396–398 future research needs, 410 as metal ion sensors colorimetric sensors, 407–409, 408, 409 fluorescence-based sensors, 404– 407, 405, 406 other sensors, 409–410 nucleic acid enzyme-metal ion interactions, 399–403 selection using in vitro evolution, 397, 397–399 double-proton transfer (DPT), 31–32, 43–46, 45, 47–48 europium Eu(III) complexes in DNA hydrolysis, 377–378, 381–382, 383–384 G-quadruplexes, role in, 59 excited state light absorbers applications in photodynamic therapy (PDT) excited state electron transfer theory, 252–253

515

excited state energy transfer theory, 254 Type I photooxidation reactions, 253–254 Type II photooxidation reactions, 254–255 groups of bimolecular interactions, 251–252 fluorescent DAPI-displacement assays, 337 gene promoters in G-quadruplex-forming sequences, 64 c-kit promotor sequence, 65, 66 c-myc-1245 sequence, 65, 66 c-myc-2345 sequence, 65, 65–66 c-myc-23456 sequence, 65, 66 glutathione reduction of Cr(VI), 466, 467 role in arsenic-induced cancers, 493, 493, 496 G-quadruplexes cation competition, 75–77 cation coordination and effect on stability, 59–61, 80–81 cation localization, 76 cation locations and coordination geometries, 69–70, 70 cation movement, 78–80 coordination of cations within d[(G4T4G4)2] crystal structures, 73–75 d(G4T4G4), 73–75 NMR studies, 75–78 coordination of cations within d[(TG4T)4], 69–73 d(TG4T), 71, 71–72 inhibition of cell proliferation, 213–215, 215 metallo-organic complexes, 215–216, 216 molecular switches, 61 numbers of bound cations, 77 Oxytricha telomere repeat analogues, 67, 67–69 platinum complexes, interaction with, 217–223, 218 promoter regions, adoption by, 64–66 structure basic, 56, 56–57 in human telomeric DNA, 61–64, 63

516

Index

loops, 58, 58–59 stoichiometry, 57, 57–58 structure of r(UG4U), 72–73 structure of telomeric DNA sequences, 213, 214 four-repeat sequences, 62–64, 63 single repeat sequences, 62 two-repeat sequences, 62 guanine. see also G-quadruplexes cis amine interactions in M-DNA adducts, 149–150 depurination, 46–47 dirhodium complexes, binding with, 303, 305 double-proton transfer, 44–46, 45 G/G crosslinks, orientations in, 140–141, 141 G-H8 resonances affected by Mn2+ adducts, 10–14, 11, 12, 13, 14 interacting sites, strength of, 6 metal ion affinities, 3 metal-DNA supramolecular chemistry 1D coordination polymers, 110– 111, 112, 114, 114, 115, 115 chelates, 124 cyclic hexanuclear platinumguanine derivative, 105–106, 106 H-bonding patterns to metals, 123, 124 square supramolecules, 102, 102– 103, 103 oxidation, 7, 469–473 platinum binding sites, possible, 138, 139–140 point defects, 48 retro models with untethered guanine bases, 143–145 rhodium(III) complexes, binding with, 307 telomeric DNA, 209, 223 highest occupied molecular orbital (HOMO), 6–7, 7 HOMO-LUMO interactions, 6, 127, 239–240, 240, 453, 454 hydrogen peroxide reduction of Cr(VI), 466, 467 intercalation, 320, 322–323 intercalator-metal conjugates, 380–382, 381, 382

ionization potentials (IP), 6, 8, 8 iridium, 100–101 iron Fe(III) complexes in DNA hydrolysis, 378–379 iron complex-peptide conjugate, 353–354 oligodeoxynucleotides adducts, 14–15 preferred coordination geometry of Fe2+ cations, 417 lanthanides, 373–374 lead DNA binding models, 34–35, 35 DNA binding parameters, 35 DNAzyme for Pb(II), 396 DNAzyme-based Pb2+ sensors, 405, 405, 407–410, 408, 409 G-quadruplexes, role in, 59 lesions. see also DNA damage; DNA repair chromium-induced, 469, 469–470, 470, 471, 472, 480 growth, 236 Lewis acidity, 301 ligands. see also nucleobases Cu(II) complexes in DNA hydrolysis, use with, 376 Fe(III) complexes in DNA hydrolysis, use with, 379 G-quadruplexes, 215, 215, 216, 218 guanine cis amine interactions, 149–150 interligand interactions, 151 lanthanide ions in DNA hydrolysis, use with, 374, 375, 377, 378 metal-DNA binding sites, role in determining, 22 metallosupramolecular-peptide conjugates, used in, 353, 353 platinum complexes, 137, 142, 142–145, 144 polyazine, used in PDT, 218, 244 ruthenium(II) probes of DNA, used in, 321–322 ancillary, 324 bridging, 325–329, 326–327, 334 flexible, 335–337, 336 intercalating, 320, 322, 323–324 solid-state structures of dynamic nucleotides, 150–151 lipophilicity, 309, 310

Index lithium G-quadruplexes coordination in d(TG4T), 71 role in, 59 lowest unoccupied molecular orbital (LUMO), 6 HOMO-LUMO interactions, 6, 127, 239–240, 240, 453, 454 magnesium basic properties of Mg2+ cations, 416–417 DNA grooves, occupancy in, 25, 25 DNAzyme for Mg(II), 396 G-quadruplexes coordination in d(TG4T), 72 role in, 59–60 polymerase catalysis Mg2+ requirement, 417–419 ion alignment, 419–420 preferred coordination geometry of Mg2+ ions, 417, 417 manganese DNA grooves, occupancy in, 25 G-H8 resonances in oligodeoxynucleotide adducts, effect on, 10–14, 11, 12, 13, 14 G-quadruplexes, role in, 59–60 Mn(II) complexes in DNA hydrolysis, 375 as relaxation probe, 9 M-DNA, 125–127, 126 M-DNA adducts internucleotide interactions, 145–147, 147, 148 solid-state structures, 150–151 mercury DNAzyme-based Hg2+ sensor, 406 metal-DNA supramolecular chemistry, 107–108, 108, 115 T-HgII-T base pair applications, 454–455 biological relevance, 455–457 crystallographic studies, 440–441, 441, 442 HgII-complex with uracil, 440–441, 441 NMR studies, 444–452, 449, 450, 451, 452 proposed structures, 440 reaction pathway, 453 research history, 439–440

517

types of HgII-T/U complexes, 444 UV, UVCD and vibrational studies, 441–444, 443 metal complex-DNA interactions. see also specific metals covalence index, correlation with, 38, 38 modes of interaction with oligonucleotides, 302 photochemical reactions, 255–257 types of DNA used in experiments, 256, 256–257 metal complex-peptide conjugates. see also specific metals binding, 347–349 bioactive peptides, 354 cleavage, 359–361 compared to metal-peptide complexes, 361 de novo designed peptides, 354–355 dipeptides, 350, 350 DNA-binding affinity, 357–358 nuclease activity on super-coiled plasmid, 360 protein fragments, 355, 355–357 sequence selectivity, 361 tripeptides glycyl-L-histidyl-L lysine (GHK), 350–352, 351, 352 Gly-Gly-Ser-CoNH2, 353, 353–354 ruthenium-peptide complexes, 351–353 metal ion-DNA interactions. see also specific metals binding, 3–4, 4, 5, 32, 38–43 DNA hydrolysis, role in, 371, 371 mechanistic investigations, importance of, 372–373 G-quadruplexes effect on stability, 59–61 locations and geometries, 69–70, 74 movement within, 78–80 role in determining structure, 61 ion reactions with hydrated electrons, 32, 32 metal-induced point defects depurination, 46–47 double proton transfer, 43–46, 45 formation, 47–48 NMR studies with oligonucleotides [Pt(dien)]+ ions, 18–21 Co2+ ions, 15

518

Index

Cu2+ ions, 15–16 Fe2+ ions, 14–15 Mn2+ ions, 10–14 Ni2+ ions, 15 Zn2+ ions, 16–18 sequence selectivity, 4 ab initio calculation studies, 6–9 photo-cleavage studies, 6, 7–9 stability constants, 34–38, 35, 36, 37 thermodynamic adsorption model of complex formation, 33–34 metal-DNA supramolecular chemistry building blocks and structural possibilities, 96–98, 97, 98 discrete architectures larger supramolecules, 105–108, 106, 107, 108 square supramolecules, 100, 102, 102–105, 103, 104, 105 triangular supramolecules, 99–102, 100, 101 H-binding patterns, 122, 122–124, 123 infinite architectures 1D coordination polymers, 109, 109–117, 110, 111, 112, 117 2D coordination polymers, 118, 118–119, 119 3D coordination polymers, 119– 121, 120, 121 potential applications, 108, 109 known structures, 95, 96 M-DNA, 125–127 metal coordination sites and base-base interactions chelates, 124 H-bonding, 121–124 metallo-DNAzymes Cu(II), 396 Mg(II), 396 nucleic acid enzyme-metal ion interactions studies fluorescence resonance energy transfer (FRET), 401–402, 402 nucleic acid enzyme-metal ion interactions, 401 nucleic acid-metal ion interactions, 401 single molecule FRET (smFRET), 402–403, 403 structural studies, 400 Pb(II), 396

metallointercalator-metallopeptide conjugates, 358–359, 359 metallonucleases, artificial, 387 metal-oxalate frameworks, 116, 117 mismatch repair, 180–182, 183, 198–199, 478–480 molecular wires, 113, 125–127 mutations metal-induced point defects depurination, 46–47 double-proton transfer (DPT), 43–46, 45 formation of point defects, 47–48 transition-type, 31, 47–48 transversion-type, 31 nickel chemical shift probe, 9 G-H8 and A-H8 resonances, effect on, 16 oligodeoxynucleotides, adducts with, 15, 16 UV spectra of various DNAs and polynucleotides, influence on, 41, 41–43, 42, 43 NMR studies cation coordination in d[(G4T4G4)2], 75–78 G-quadruplexes cation coordination and effect on stability, 59–61 cation movement, 78–80 coordination of cations within d[(G4T4G4)2], 75–78 human telomeric sequences, structure of, 62–63, 63 loops, 58–59 Oxytricha telomeric sequences, structure of, 67, 67–69 sequence selectivity in DNA-metal binding, 9–21 methodology, 9–10 model systems, 10 oligodeoxynucleotides-transition metal adducts, 10–21 structure of the T-HgII-T base pair 15 N-HgII-15N covalent linking, 450, 450, 451 J-coupling values, 451 nucleoside-metal ion systems, 444–448

Index oligonucleotide-metal systems, 448–449, 449 T-HgII-T base pair, 449–452 nucleases, artificial mechanistic pathways, 361–362 metallonucleases, 387 research aims, 359 nucleic acid enzyme catalysis catalytic centres, types of, 418 Mg2+ ions, requirement for two DNA and RNA polymerases, 417–419 RNase H and MutH, 420–427, 421 one-metal-ion mechanism, 428–429 two-metal-ion mechanism advantages, 428, 429 catalytic specificity enhancement, 423–424 effect of substrate on binding of metal ions, 422–423 ion alignment, 419–420, 424, 427 Mg2+ ions, 417–419, 420–422 movement of two metal ions, 425 prediction, 429–430 separate and joint functions of metal ions, 426–427, 427 types of catalytic centres, 418–419 nucleobases. see also adenine; cytosine; guanine; thymine; uracil as bridging ligands 1D coordination polymers, 109, 110, 110–113 3D coordination polymers, 119– 121, 120 as bridging ligands with anions 1D coordination polymers, 114–117 2D coordination polymers, 118–119 Cr(VI)-induced damage and repair base excision repair (BER) pathway, 473 lesion pathways, 468–473, 469, 470, 471, 472 reduction potentials, 469 formation of chromium-DNA adducts, 477–478, 478 metal-DNA supramolecular chemistry building blocks, 96–97, 97 chelating, 124, 125 self-pairing, 122, 122–124, 123 preferred metal ion binding sites, 3

519

nucleotide excision repair, 178–180, 181, 193–198, 481–482, 497–498 oligonucleotides formation of G-quadruplexes, 57 metal complexes conjugated with, 384 modes of interactions with, 302 NMR studies with metal complexes, 448–449, 449 NMR studies with metal ions [Pt(dien)]+ ions, 18–21 Co2+ ions, 15 Cu2+ ions, 15–16 Fe2+ ions, 14–15 Mn2+ ions, 10–14 Ni2+ ions, 15 Zn2+ ions, 16–18 platinum complexes, interaction with, 160–163, 161, 162 oligonucleotides, platinated crosslinking, 276–278, 277, 278 duplex, 289–294, 290, 291, 293 triplex, 294–295 drug requirements, 276 sulfur-platinum-nitrogen adducts, 286–287, 287 synthesis base-modified oligonucleotides, 286–289, 289 partially protected oligonucleotides, 283–284, 284 platination site, identification of, 288 protected oligonucleotides, 285, 285–286, 286 solid-phase synthesis, 281–283, 282 unmodified oligonucleotides, 279–281, 280 unprotected oligonucleotides, 281, 281 therapeutic applications antisense oligonucleotides, 273– 274, 274 aptamers, 275–276 microRNA targeting, 275 ribozymes, 274–275 RNA interference, 275 triplex-forming oligonucleotides, 275

520

Index

organotin industrial uses, 308 lipophilicity and toxicity, correlation between, 309 organotin complex-DNA interactions, 309–311, 310 toxicity, 308 osmium polyazine complexes used in PDT electronic excitation and unimolecular decay, 242, 243, 247–248, 248 phototoxicity, 265–266 oxaliplatin, 135–136, 136, 155, 155–156, 188 p53 tumour suppressor protein, 496, 498 palladium, 103, 103 paramagnetic relaxation, 9–10 peptides. see metal complex-peptide conjugates; metallointercalatormetallopeptide conjugates peraklyammonium groups, 383–384 pH effect on H-conformer preferences in platinum complexes, 149, 150 influence on organotin-DNA interactions, 310 phosphoryl transfer reactions, 415–416, 416 photo-cleavage studies, 6, 7–8, 8 photodynamic therapy (PDT) cell studies with metal complexes cellular uptake, 263–265 complexes used, 263 phototoxicity, 265, 265–266 clinical treatments, 236–237, 237 definition, 236 designing polyazines for DNA photomodification design considerations, 238, 257 ground state interactions, 257–258 photochemical degradation mechanisms, 258–262 DNA type used, influence of, 256, 256–257 PDT agents, 237–238 research needed, 267 rhodium complexes, 305–306, 306 photophysical processes of interest in PDT research

bimolecular excited state interactions, 251–255 electronic excitation, 239–240, 240, 241 unimolecular electronic excited state decay, 240–242, 242 picoplatin, 136, 136 plasmid DNA, 371–372, 372 platination DNA, effects on, 140 enhancement of biological effects, 276–278, 278 oligonucleotides base-modified, 286–289 duplex crosslinking, 289–294 partially protected, 283–284, 284 protected, 285, 285–286, 286 solid-phase synthesis, 281–283, 282 triplex crosslinking, 294–295 unmodified, 279–281, 280 unprotected, 281, 281 telomeric DNA, 225 platinum. see also cisplatin (and related platinum complexes); platination; platinum drugs; transplatin binding to DNA, 138, 138–140 crosslinking DNA duplexes, use in, 289–294, 290 DNA damage and repair, 176–177 metal-DNA supramolecular chemistry 1D coordination polymers, 109– 110, 110, 111 2D coordination polymers, 118, 118 large structures, 105–107, 106, 107, 108 metallocalixarenes, 104, 105, 105, 105, 105 supramolecular triangles and squares, 100, 101, 101–102 oligodeoxynucleotides, adducts with, 18–21, 19, 20, 21 peptide-tethered complexes, 350, 350 telomerase, interaction with, 223–225 telomeric DNA duplexes, interaction with, 223 platinum drugs. see also cisplatin (and related platinum complexes) activity series, 137 delivery to tumours, 136 design factors to consider, 200 drugs in use, 135–136, 136

Index effects of DNA repair, 177 repair of DNA damage base excision repair, 193 homologous recombination repair, 199–200 mismatch repair, 198–199 nonhomologous end-joining, 199 nucleotide excision repair, 193–198 structure-activity relationships (SARs), 137 types, 135–136, 136 polyazine complexes, transition metal cellular uptake, 263–265 common electronic transitions, 243 complexes used in PDT, 263 electronic excitation and unimolecular decay osmium complexes, 242, 243, 247– 248, 248 rhodium complexes, 242, 248–250, 249, 250 ruthenium complexes, 242, 243– 247, 247, 248 ruthenium-rhodium mixed complexes, 250–251, 251 ligands, 244 photophysical properties, 245–246 toxicity studies, 265–266 potassium G-quadruplexes competition with Na+ , 77 coordination in d[(G4T4G4)]2, 74 coordination in d(G4T4G4), 73–74 human telomeric sequences, 63, 63–64 melting temperatures, 60 residence times, 78 role in, 59, 61 praseodymium, 377 proteins repair of DNA modified by antitumour platinum compounds AAG, 189 base excision repair, 189 DNA-PK, 192 high-mobility group proteins, 193 hMutLα, 191–192 hMutSα, 191–192 homologous recombination repair, 192 mismatch repair, 191–192

521

nonhomologous end-joining, 192 nucleotide excision repair, 189–191 RNA polymerase II, 191 RPA, 190, 191 translesion synthesis DNA polymerases, 192–193 XPA, 190 XPC-HR23B, 190–191 in telomeres, 210–211, 211 proton tunneling, 44–46, 45 purines metal-DNA supramolecular chemistry cyclic trimeric complexes, 100–101 large structures, 105–106 molecular squares, 100, 102, 102– 103, 104 molecular triangles, 99–101, 100, 101 tetrameric complexes, 100–101 structure, 97 pyrimidines metal-DNA supramolecular chemistry large structures, 106–108 metallocalixarenes, 103–105 molecular triangles, 101–102 structure, 97 reactive nitrogen species, 495–496 reactive oxygen species, 495–496 recombination repair, 482 retro modelling cisplatin analogues, 141–143, 142 untethered guanine bases, 143, 143–145, 144 rhodium complexes used in PDT, 305–306, 306 dirhodium (II) complexes, 303–305, 304 intercalator-metal conjugates, 381, 381 metal-DNA supramolecular chemistry, 100–101, 113, 114 polyazine complexes used in PDT cellular uptake, 264 DNA photochemical degradation, 259–260 electronic excitation and unimolecular decay, 242, 248– 251, 249, 250, 251 phototoxicity, 265, 265–266 rhodium (I) complexes, 302–303 rhodium (III) complexes, 306–307

522

Index

rhodium complex-peptide conjugate, 354, 358–359, 359 site-specific rhodium-DNA interactions, 307, 307–308 ribozymes, 274–275, 400 RNA dinuclear ruthenium, interaction with, 334–335 Okazaki fragments, 209–210, 210 RNA cleavage, 415–416, 416, 426–427, 427 RNA interference, 275 RNA polymerases, 418, 418–419 RNA repair, 177 RNA synthesis, 415, 416 rubidium, 23, 23, 59 ruthenium advantages of, as DNA probe, 319–320 dinuclear complexes as DNA probes advantages over mononuclear species, 325 affinity chromatography, 337–338 applications, potential, 338–339 binding behavior, 325–329 DNA bulge sequences, interaction with, 330–332, 331, 333 DNA hairpin sequences, interaction with, 332–334 duplex DNA, interaction with, 329, 329–330 flexible ligands, 335–337, 336 fluorescent DAPI-displacement assays, 337 research needs, 338 RNA sequences, interaction with, 334–335 S-O-S bridge, 325, 327 stereoisomers, 329 intercalator-metal conjugates, 381, 381 metal-DNA supramolecular chemistry, 100–101, 101 mononuclear complexes as DNA probes binding site and orientation, 322–324 enantioselectivity, 320 influence of nature of ligands on binding, 323–324 intercalation, 320, 322 polyazine complexes used in PDT cellular uptake, 264

electronic excitation and unimolecular decay, 242, 243– 247, 247, 248, 250–251, 251 photobinding of DNA, 259, 260 phototoxicity, 265, 265–266 ruthenium complex-peptide conjugate de novo designed peptides, 354–355 protein fragments, 355–357 satraplatin, 136, 136 shelterin (telosome), 210–211, 211 silver metal-DNA supramolecular chemistry 1D coordination polymers, 109– 110, 110, 111 2D coordination polymers, 118– 119, 119 3D coordination polymers, 120– 121, 121 sodium DNA grooves, occupancy in, 24 effect on stability constants of metal ions, 36, 36–37 G-quadruplexes competition with K+, 77 competition with NH4+, 75–76 coordination in d[(G4T4G4)]2, 74 coordination in d[(TG4T)4], 70 coordination in d(G4T4G4), 73 coordination in d(TG4T), 71, 71, 72 localization, 70, 76 melting temperatures, 60 residence times, 78–79 role in, 59, 61 spine of hydration, 23, 24 stability constants of metal-DNA complexes covalence index, correlation with, 38, 38 determination, 34–36, 35, 36 DNA G-C content, effect of, 37, 37–38 ionic strength, effects of, 36, 36–37 strontium G-quadruplexes coordination in r(UG4U), 72–73 role in, 59, 60 sugar oxidation, Cr(VI)-induced, 474–477, 475, 476 tautomeric transformations, 44, 47 telomerase, 212, 212–213, 223–225 telomeres, 209–211, 210

Index telomeric DNA G-quadruplexes, 213, 214 human, 61–64, 63 Oxytricha, 67, 67–69 platinum complexes, interaction with, 223, 225 telosome (shelterin), 210–211, 211 thallium G-quadruplexes coordination in d[(G4T4G4)]2, 74 coordination in d(G4T4G4), 75 coordination in d(TG4T), 71, 72 localization, 76–77 residence times, 78 role in, 59 thermodynamic adsorption model of complex formation, 33–34 thymine loop coordination in G-quartets d(G3T4G3), 68 d(G3T4G3C), 61 d(G3T4G4), 69 d(G4T4G3), 68–69 d(G4T4G4), 67–68, 73–75 metal ion affinities, 3 metal-DNA supramolecular chemistry 1D coordination polymers, 109 chelates, 124, 125 H-bonding patterns to metals, 122 large supramolecules, 106–107, 107 platinum binding sites, possible, 138 thymine-thymine base pair, HgII-mediated applications, 454–455 biological relevance, 455–457 structural studies crystallographic studies, 440–441, 441, 442 HgII-complex with uracil, 440–441, 441 NMR studies, 444–452, 449, 450, 451, 452 proposed structures, 440 reaction pathway, 453 research history, 439–440 types of HgII-T/U complexes, 444 UV, UVCD and vibrational (IR/Raman) spectral studies, 441–444, 443

523

tin organotin industrial uses, 308 lipophilicity and toxicity, correlation between, 309 organotin complexes-DNA interactions, 309–311, 310 toxicity, 308 transplatin, 136, 136, 140 platinated oligonucleotides crosslinking experiments, 279–280, 280, 292–295, 293 isomerization reaction, 290–291, 291 regioselective platination, 286–288 uracil HgII-complex, 440–441, 441 metal-DNA supramolecular chemistry 1D coordination polymers, 109, 109–110 chelates, 124 H-bonding patterns to metals, 122, 124 large supramolecules, 106–107, 107 square supramolecules, 105, 105 uranium, 405–406, 406 UV studies HgII-nucleobase complexes, 441 study of metal-DNA interactions bathochromic shift, 39, 39–41, 40 double-proton transfer (DPT) measurements, 43–46 influence of GC composition, 41–43 influence of Ni2+ on spectra of DNAs and polymerases, 41, 41–43, 42, 43 vibrational studies with HgII-nucleobase complexes, 441–444, 443 X-ray crystallography studies cation-groove interaction studies, 22 G-quadruplexes coordination of cations within d[(G4T4G4)2], 73–75 coordination of cations within d[(TG4T)4], 69–73 metal ion–ribozyme complexes, 400

524

Index

zinc DNA cleavage, 358 G-quadruplexes, role in, 59–60 oligodeoxynucleotides, adducts with, 16–18 chemical shift variation for G-H8 resonances, 16–17, 17 effect of ZnCl2 concentrations, 17 preferred coordination geometry of Zn2+ cations, 417, 417

zinc-finger peptides in DNA hydrolysis, 384 Zn(II) complexes in DNA hydrolysis, 374–375, 375, 379–380, 380, 383 intercalator-metal conjugates, 381, 382, 383 Zn(II) complexes in DNA oxidative cleavage, 373, 373 zirconium, 384–385